Sarcomeres are the basic contractile units within cardiac myocytes, and the collective shortening of sarcomeres aligned along myofibrils generates the force driving the heartbeat. The alignment of the individual sarcomeres is important for proper force generation, and misaligned sarcomeres are associated with diseases, including cardiomyopathies and COVID-19. The actin bundling protein, α-actinin-2, localizes to the ‘Z-Bodies” of sarcomere precursors and the ‘Z-Lines’ of sarcomeres, and has been used previously to assess sarcomere assembly and maintenance. Previous measurements of α-actinin-2 organization have been largely accomplished manually, which is time-consuming and has hampered research progress. Here, we introduce sarcApp, an image analysis tool that quantifies several components of the cardiac sarcomere and their alignment in muscle cells and tissue. We first developed sarcApp to utilize deep learning-based segmentation and real space quantification to measure α-actinin-2 structures and determine the organization of both precursors and sarcomeres/myofibrils. We then expanded sarcApp to analyze ‘M-Lines’ using the localization of myomesin and a protein that connects the Z-Lines to the M-Line (titin). sarcApp produces 33 distinct measurements per cell and 24 per myofibril that allow for precise quantification of changes in sarcomeres, myofibrils, and their precursors. We validated this system with perturbations to sarcomere assembly. We found perturbations that affected Z-Lines and M-Lines differently, suggesting that they may be regulated independently during sarcomere assembly.
cardiac myocytesarcomeremyofibrilResearch organismHumanhttp://dx.doi.org/10.13039/100000057National Institute of General Medical SciencesR35 GM125028BurnetteDylan Tylerhttp://dx.doi.org/10.13039/100009633Eunice Kennedy Shriver National Institute of Child Health & Human DevelopmentT32 5T32HD007502Neininger-CastroAbigail CSanchezZachary Chttp://dx.doi.org/10.13039/100000057National Institute of General Medical SciencesT32 5T32GM008320HayesJames Bhttp://dx.doi.org/10.13039/100000968American Heart Association836090HayesJames Bhttp://dx.doi.org/10.13039/100000968American Heart Association1070985SanchezZachary Chttp://dx.doi.org/10.13039/100000050National Heart, Lung, and Blood InstituteF31 HL136081FenixAidan Mhttp://dx.doi.org/10.13039/100000968American Heart Association18PRE33960551TanejaNilayThe funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.Author impact statementSoftware is introduced for the rapid and comprehensive quantification of sarcomere-containing myofibrils and their precursors in cardiac myocytes, aiding in the advancement of cardiovascular cell biology investigations.publishing-routeprcIntroduction
The sarcomere is the fundamental unit of contraction in striated muscle (Au, 2004). Sarcomeres individually contract to generate a force, and sarcomeres organize and align within myofibrils to augment that force. These forces enable skeletal muscle to coordinate organism-level locomotion and cardiac muscle to drive the heartbeat (Hersch et al., 2013; Wang et al., 2021). Sarcomere structure and alignment is critical to muscle function, and dysfunctional sarcomeres and myofibrils have been implicated as causes of disease (Eschenhagen et al., 2015; Harvey and Leinwand, 2011; Olsson et al., 2004).
Since the mid-20th century, the question of how sarcomeres are assembled and subsequently organized into higher-order myofibrils has been a much-debated topic, with multiple separate lines of research emerging to support a small host of potential models (Dlugosz et al., 1984; Sanger et al., 1984; McKenna et al., 1985). Generally, the models of sarcomere assembly that are backed by the most data posit that sarcomeres arise from either (i) a ‘parts-wise’ assembly, where precursors of unique sarcomeric regions (e.g., the thin [actin] or thick [myosin] filament) are first assembled separately, then ‘stitched’ together to generate the final structure (i.e., Stitching Model) or (ii) a more direct assembly, where a precursor forms that resembles a sarcomere but has a subset of non-sarcomeric proteins that are later replaced by sarcomeric proteins (i.e., Pre-myofibril Model) (Dlugosz et al., 1984; Rhee et al., 1994). Recently, our group has shown that sarcomeres descend directly from muscle stress fibers (MSFs), with our data supporting a subset of predictions made by both models, but not necessarily favoring either model (Fenix et al., 2018). All models of sarcomere assembly are currently limited in their testability because the field lacks a tractable workflow for high-throughput analysis.
Recently, our lab has shown that human induced pluripotent stem cell-derived cardiac myocytes (hiCMs) can be harnessed as a tractable model for routine imaging of sarcomere assembly in live CMs (Fenix et al., 2018; Fenix et al., 2016). In our assay, hiCMs assemble sarcomeres de novo, and directly from MSFs, within 24 hr. Despite having powerful tools for imaging, our analysis workflow is still hampered by technical bottlenecks related to (i) image binarization and (ii) manual quantification. Binarization via classical methods relying on image pixel intensity (e.g., Otsu’s method) is often performed manually – slowly – and can become muddled by a multitude of factors, including (i) high-intensity artifacts related to nonspecific localization of antibodies, (ii) low-intensity staining due to high background levels/noise, and/or (iii) multi-valent interactions by the target protein itself occluding the specific structure of interest (Otsu, 1979; Sankur, 2004; Uchida, 2013). Meanwhile, we find that even well-binarized, high-resolution images can require several hours of manual quantification per image.
Deep learning has emerged as a powerful potential solution for rapid, accurate binarization of complex grayscale images. Deep learning-based binarization can be accomplished using a framework called a U-Net (Ronneberger and Brox, 2015). A U-Net trains a deep learning model to convert images into only background (noise) and foreground (signal) by matching images to expert-annotated ‘ground truth’ binaries – that is, images manually annotated by the user. Compared to manually performed binarization, U-Net offers the advantages of superior accuracy and speed, but at a high upfront cost, since implementation of a U-Net requires a user with advanced background in mathematics, experience with machine learning, and proficiency in writing software code (Ronneberger and Brox, 2015; Yin et al., 2022).
While a sarcomere contains 100+ proteins, the field has often resorted to ‘proxying’ sarcomere assembly by staining, often exclusively, for α-actinin-2 (Fenix et al., 2018; Henderson et al., 2017; Chopra et al., 2018; Ehler et al., 1999; Hinson et al., 2015; Taneja et al., 2020). α-actinin-2 marks the sarcomeric Z-Lines, which border the sarcomeric contractile machinery (Murphy and Young, 2015; Ribeiro et al., 2014). In the last decade, several methods for automatic quantification of Z-Lines have emerged, with the most widely used methods relying on calculations made in frequency space (i.e., by using a Fourier transformation of the image) (Chopra et al., 2018; Hinson et al., 2015). While frequency space calculations can detect repeating patterns created by adjacent Z-Lines, current methods yield no information about Z-Lines in real space (e.g., number, size, spacing, or organization within myofibrils), about sarcomere precursors, or about any sarcomeric components other than Z-Lines. Moreover, it is not yet clear quite how the assembly of Z-Lines themselves relates either spatially or temporally to the assembly of other sarcomere components, or vice versa.
Here, we present a high-throughput, automated, and non-biased quantification scheme for sarcomere assembly. Our approach is two-pronged and includes a unique method for both (1) rapid, accurate binarization of images that can consistently separate true signal from noise and structures of interest from those that are not of interest and (2) automated quantification of relevant sarcomeric and pre-sarcomeric structures within the binarized images. Towards (1), we have developed ‘yoU-Net,’ a U-Net-based framework with a user-friendly graphical user interface (GUI) that enables a user with little to no experience to binarize images with the power and flexibility afforded by deep learning. We demonstrate that yoU-Net can accurately binarize images of CM sarcomeres and sarcomere precursors using multiple unique sarcomere stains. Towards (2), we have developed ‘sarcApp,’ a software that automatically annotates and calculates 50+ unique descriptive outputs of sarcomeres and sarcomere precursors in real space using binarized images. Using yoU-Net and sarcApp, we demonstrate that some perturbations to sarcomere assembly preferentially affect specific sarcomere components (e.g., M-Lines) more than others (e.g., Z-Lines). Our data emphasize the need for more high-throughput studies into sarcomere assembly that includes multiple sarcomere components, and altogether point to a disjointed or modular sarcomere assembly program within CMs that is more nuanced than was previously believed. The tools presented herein will facilitate future high-throughput studies to develop a more mature and complex model of sarcomere assembly.
ResultsyoU-Net enables user-friendly binarization of dynamic grayscale images
Our overarching goal was to develop a method for fast and accurate quantification of sarcomeres within hiCMs using our sarcomere assembly assay. We faced two primary bottlenecks in our workflow, the first of which was a slow and cumbersome image binarization process. Binarization facilitates image quantification by converting the complex, raw image of a biological specimen into a far simpler image containing only background (non-relevant information) and signal (relevant information). A well-binarized image should retain high fidelity to the original image but contain only the relevant structures of interest superimposed over a blank background. Individual pixels within a binary image are stripped of dynamic information present within the raw image and are re-assigned to represent only either background or signal – binary images are therefore often represented in black (background) and white (signal).
Traditional methods of binarization fall short for images of sarcomeres and sarcomere precursors in hiCMs due to the dynamic range of pixel intensities present within the raw images. In images of hiCMs 24 hr after plating, sarcomere precursors and sarcomeres are each stained by α-actinin-2 (Figure 1A), but the Z-Lines of the sarcomeres are markedly brighter than the Z-Bodies of the precursor structures (i.e., MSFs). Classical, intensity-based binarization methods force a user to make a choice in this scenario – specifically, whether to accurately binarize based on gray levels of the sarcomere precursors or of the sarcomeres themselves (without the consistent option for both). Classical Otsu-based thresholding (using FIJI) of a representative cell in Figure 1A shows that binarizing this cell according to Z-Lines oversaturates Z-Bodies into an inseparable clump (Figure 1A, arrowhead), leaving a user unable to accurately quantify both structures from this single binary.
yoU-Net follows a U-Net architecture to binarize immunofluorescence images.
(A) Representative images of α-actinin-2 and actin filaments (phalloidin) in a human induced pluripotent stem cell-derived cardiac myocyte (hiCM). The α-actinin-2 binary was thresholded in FIJI using Otsu’s method. Orange arrowhead denotes Z-Bodies. (B) yoU-Net architecture from input image to output binary. Details can be found in the Methods and in Figure 1—figure supplement 1. (C) yoU-Net-generated binary of α-actinin-2, predicted using the trained U-Net described in (B). Orange arrowhead: Z-Bodies. (D) Model of muscle stress fibers (MSFs) and myofibrils during sarcomere formation. Black arrow denotes direction of MSF translocation as α-actinin-2-positive Z-Bodies elongate and coalesce to form Z-Lines.
Details of yoU-Net architecture and trained model generation.
(A) U-Net architecture details used for this manuscript (details can be adjusted in yoU-Net). (B) Dice coefficient formula, used for validating trained models. (C) Representative fluorescence image and matched annotated binary of a human induced pluripotent stem cell-derived cardiac myocyte (hiCM) with α-actinin-2 localized. (D) Representative fluorescence image and matched annotated binary of an hiCM with titin localized. (E) Representative fluorescence image and matched annotated binary of an hiCM with myomesin localized.
Our lab was well-positioned to approach this problem by constructing a deep learning-based model called a U-Net (Ronneberger and Brox, 2015). A U-Net enables a user to train a deep learning model to automatically binarize images with high precision and speed. Model ‘training’ involves matching raw images to user-annotated binaries – called ‘ground truth’ binaries – using a complex set of mathematical convolution/downsampling and deconvolution/upsampling steps. The architectural layout of a U-Net is represented by Figure 1B.
We constructed our own U-Net and trained it using 1000 training steps (100 epochs/steps of 10 iterations each) of hiCMs stained for α-actinin-2. After training, our U-Net can reliably and automatically predict high-fidelity binaries of hiCM α-actinin-2 stains that are resistant to artifacts related to differences in pixel intensity and can accurately separate individual Z-Bodies (Figure 1C, arrowhead). Recognizing that other labs would benefit from the flexibility and accuracy of our U-Net, we also wrote software encoding for a GUI to accompany our U-Net in the hopes of making it more user-friendly and broadly accessible. Our U-Net and GUI together are known as ‘yoU-Net’ and are available, open source with details in the supplement (Figure 1—figure supplement 1). We used yoU-Net to binarize images throughout the remainder of this study.
sarcApp quantifies muscle stress fibers, sarcomeres, and myofibrils
The next bottleneck within our workflow was the issue of manual quantification, which in some cases can require several hours for a single CM. A meaningful quantitative output of sarcomere assembly would discriminate between sarcomeres and sarcomere precursors (MSFs), and produce readouts for each. Sarcomere assembly involves the formation MSFs near the cell edge that give rise to sarcomeres typically closer to the cell center (Figure 1D). Within MSFs are punctate α-actinin-2-containing structures termed ‘Z-Bodies,’ which gradually elongate/concatenate over time and eventually transition into the Z-Lines of sarcomeres (Figure 1D). While the Z-Body to Z-Line transition exists along a continuum, it was necessary for us to define a binary transition point on the basis of α-actinin-2 structure length when a Z-Body ‘becomes’ a Z-Line. We tasked individual lab members with independently annotating 10 images each to determine which structures constituted a Z-Body and which were Z-Lines. Together, we defined a potential Z-Line to be any α-actinin-2-containing structure with a length >1.4 µm, with all other structures being potential Z-Bodies. Because many organisms have variable sarcomere spacing and structure, this variable along with all others discussed (lengths, minimum number of structures, etc.) are customizable and can be easily redefined by the user for their unique model system (Reedy and Beall, 1993).
Using our predefined size criterion for Z-Lines and Z-Bodies, we next desired a method capable of automated detection, measurement, and descriptive output of both structures. Towards this goal, we developed sarcApp, a software-based code that classifies each α-actinin-2 structure within a binary image as a Z-Line (Figure 2A and B) or Z-Body (Figure 2C and D) and further assigns each to either a myofibril (Figure 2B) or MSF (Figure 2D). All structures identified as potential Z-Lines are paired to other potential Z-Lines in the cell based on shape, orientation, and location, then assigned to myofibril based on orientation to other Z-Lines (Figure 2A and B). Z-Lines, which belong to a myofibril, are considered ‘confirmed.’ This eliminates singular α-actinin-2-positive adhesions. Next, each potential Z-Body is paired to other Z-Bodies based on shape and location, then assigned to an MSF based on orientation to other nearby Z-Bodies (Figure 2C and D). Like with Z-Lines, only Z-Bodies that belong to an MSF are considered ‘confirmed.’ After detecting, annotating/assigning, and measuring all potential Z-Lines and Z-Bodies within a given cell, sarcApp automatically compiles and generates three spreadsheets summarizing the data on a per-cell, per-myofibril, and per-MSF basis. Cell spreadsheets generated include myofibrils per cell (Figure 2E), Z-Lines per cell (Figure 2F), Z-Line lengths (Figure 2G), myofibril persistence lengths (Figure 2H), MSFs per cell (Figure 2I), Z-Bodies per cell (Figure 2J), Z-Body lengths (Figure 2K), and MSF persistence lengths (Figure 2L). Myofibril spreadsheets include analysis of myofibril organization within the cell by calculating the angle of each myofibril relative to the nearest cell edge (Figure 2M–O), as well as number of Z-Lines, average Z-Line spacing, persistence length, absolute angle of the myofibril long axis (considering the image edges as the X and Y axes), average Z-Line length, distance from the center point of the myofibril to the nearest parallel cell edge, absolute angle of the nearest cell edge, and relative angle of the myofibril to the edge (a measure of parallelism); all on a per individual myofibril basis. MSF spreadsheets include number of Z-Bodies, average Z-Body spacing, and persistence length for each MSF.
Quantifying sarcomere and myofibril organization using α-actinin-2 binaries.
(A) Z-Lines and myofibrils identified by sarcApp. Each line denotes a Z-Line, and each different color represents a different myofibril. (B) Quantification scheme for myofibrils and Z-Lines. Details can be found in Figure 2—figure supplement 1. (C) Z-Bodies identified by sarcApp. Each red circle denotes a Z-Body. (D) Quantification scheme for muscle stress fibers (MSFs) and Z-Bodies. Details can be found in Figure 2—figure supplement 1. (E) Distribution of myofibrils per human induced pluripotent stem cell-derived cardiac myocyte (hiCM) plated for 24 hr (N = 188 cells; four biological replicates). (F) Distributions of Z-Lines per hiCMs from (E). (G) Distribution of average Z-Line length per cell from (F) (N = 104 cells). (H) Distribution of average myofibril persistence lengths per cell from (G) (N = 104 cells; quantification details can be found in Figure 2—figure supplement 1). (I) Distribution of MSFs per cell from (E). (J) Distribution of Z-Bodies per cell from (E). (K) Distribution of average Z-Body length per cell from (E). (L) Distribution of average MSF persistence lengths per cell from (E). (M) Myofibril long axes identified by sarcApp. (N) Quantification scheme for myofibril angle relative to edge. Briefly, the closest edge segment to the myofibril long axis (perpendicularly) is used as the reference angle, and the numerical output is the difference between the myofibril long axis angle and the reference edge angle. (O) Distribution of myofibril orientation relative to cell edge in the same cells as (E) (N = 188 cells, 1217 myofibrils). Note that most myofibrils are relatively parallel to the edge in hiCMs plated for 24 hr.
Details of geometric calculations used in sarcApp.
(A) How Z-Line spacing is calculated in sarcApp. In short, for each pair of Z-Lines, a line is measured from the centroid of one Z-Line to the next Z-Line using a line drawn perpendicularly to the first Z-Line. The same is done vice versa: a line from the second Z-Line to the first is measured. The averaged perpendicular distances comprise the ‘true’ distance. (B) How myofibril persistence length is calculated. In short, persistence length is the sum of all true Z-Line distances across an entire myofibril. If Z-Lines are longitudinally split, the Z-Line distances are averaged. (C) How laterally linked Z-Lines (H structures) are solved in sarcApp. In short, the length, width, and centroid of the H structure are used to back calculate the predicted centroid of each individual Z-Line.
sarcApp must take certain liberties to parameterize and predict MSF/myofibrillar structures within CMs. For example, Z-Lines are assigned to individual myofibrils in part by inter-Z-Line distance (which we define as being <3 µm apart using centroid-to-centroid distance) but also by Z-Line long-axis angle (which we define as being within 30°, accounting for cell curvature). These measurements are customizable as some species have varied inter-Z-Line distance or varied cell curvature. We increase the computational speed of sarcApp by measuring Z-Line spacing using the distance between each Z-Line’s centroid as opposed to parallel spacing, which can produce slightly different results (Figure 2—figure supplement 1A). sarcApp defines a myofibril as a structure with a linear collection of four or more Z-Lines (i.e., three sarcomeres). Potential Z-Lines that are not assigned to a myofibril are not quantified, as they could represent adhesions, which are also α-actinin-2-positive (Taneja et al., 2020). Any structure with a long axis <1.4 µm is defined as a potential Z-Body. Z-Bodies are assigned to MSFs with the criteria that the Z-Bodies are (1) <3 µm apart and (2) have centroids that form a line within 30° parallel to the edge, indicative of a typical Z-Body organization with MSF actin filaments (Fenix et al., 2018).
Blebbistatin treatment reduces Z-Line assembly in hiCMs
To validate the functionality and accuracy of sarcApp, we paired our lab’s sarcomere assembly assay (Fenix et al., 2018; Taneja et al., 2020) with a well-established inhibitor of sarcomere assembly: Blebbistatin, a pan-myosin II inhibitor (Limouze et al., 2004). In brief, we trypsinized and replated hiCMs. During this process, the hiCMs lose and reform their sarcomeres, allowing us to monitor sarcomere assembly. Because this assembly begins at the cell edge and moves rearward toward the cell center, a snapshot of a hiCM after 24 hr of spreading (once sarcomeres have begun to form and myofibrils begin to align) contains information on early, mid, and late sarcomere formation at the leading edge, behind the edge in the lamella, and toward the center of the cell, respectively (Fenix et al., 2018).
Blebbistatin inhibits myosin II-based contractility and Blebbistatin-treated hiCMs exhibit reduced Z-Line assembly in our assay (Figure 3A, Figure 3—figure supplement 1), consistent with other reports (Chopra et al., 2018; Liu et al., 2016). Platinum-replica electron microscopy (EM) shows that Blebbistatin-treated hiCMs do not appear to have well-defined Z-Lines and have a disordered overall orientation of MSFs/myofibrils compared to controls (Figure 3B). As measured by sarcApp, hiCMs treated with Blebbistatin exhibited fewer myofibrils per cell (Figure 3C), fewer Z-Lines per cell (Figure 3D), decreased myofibril persistence length (Figure 3E), decreased Z-Line length (Figure 3F), and decreased overall size of all α-actinin-2-positive puncta/structures (Figure 3G). sarcApp also reported that, while the myofibrils of control hiCMs are oriented approximately parallel to the nearest cell edge (Figure 3H; similar to Figure 2M–O), the myofibrils of Blebbistatin-treated hiCMs are more randomly oriented (Figure 3H), suggesting a disorganized assembly process. Altogether, these data generated by sarcApp are consistent with a wide range of previous studies showing that Blebbistatin inhibits sarcomere assembly (Chopra et al., 2018; Liu et al., 2016).
Blebbistatin treatment ablates Z-Line formation in human induced pluripotent stem cell-derived cardiac myocytes (hiCMs).
(A) Representative images and insets of α-actinin-2 and F-actin in hiCMs treated with DMSO, 50 µM Blebbistatin, or 100 µM Blebbistatin. (B) Representative platinum replica EM image of a control hiCM and a 50 µM Blebbistatin-treated hiCM. Arrows indicate an elongated Z-Line in the DMSO-treated hiCM and a Z-Body in the Blebbistatin-treated hiCM. (C) Myofibrils per cell in hiCMs (N = 118 DMSO cells, 108 50 µM Blebbistatin cells, and 93 100 µM Blebbistatin cells; four biological replicates). (D) Z-Lines per cell in hiCMs from (C). (E) Average myofibril persistence length per cell in hiCMs from (C) (N = 104 DMSO control cells, 45 50 µM Blebbistatin cells, and 45 100 µM Blebbistatin cells). (F) Average Z-Line length per cell from (E). (G) Average size of all α-actinin-2-positive puncta in hiCMs from (C). (H) Myofibril orientation relative to the cell edge segment closest to the myofibril center, perpendicularly. N = 4 biological replicates, 1217 DMSO control myofibrils, 385 50 µM Blebbistatin myofibrils, 220 100 µM Blebbistatin myofibrils. Blebbistatin treatment for 6 and 12 hr can be found in Figure 3—figure supplement 2 and Figure 3—figure supplement 3.
α-actinin-2 quantification and organization in Blebbistatin-treated human induced pluripotent stem cell-derived cardiac myocytes (hiCMs) 6 hr post-plating.
(A) Myofibrils per cell in cells treated with DMSO or 50/100 µM Blebbistatin. N = 3 biological replicates, 73 control cells, 54 50 µM Blebbistatin cells, and 72 100 µM Blebbistatin cells. Myofibrils defined as having four or more Z-Lines in a row. Details in Figure 2—figure supplement 1. (B) Z-Lines per cell in hiCMs from (A). (C) Average myofibril persistence length per cell in hiCMs from (B). N = 59 control cells, 34 50 µM Blebbistatin cells, and 51 100 µM Blebbistatin cells (only cells with myofibrils were quantified for C–E). (D) Average Z-Line length per cell in hiCMs from (C). (E) Average spacing between Z-Lines per cell in hiCMs from (C). (F) Muscle stress fibers (MSFs) per cell in hiCMs from (A). (G) Z-Bodies per cell in hiCMs from (A). (H) Average MSF persistence length per cell in hiCMs from (A). N = 3 biological replicates, 73 control cells, 54 50 µM Blebbistatin cells, and 72 100 µM Blebbistatin cells (only cells with MSFs were quantified for H–J). (I) Average Z-Body length per cell in hiCMs from (H). (J) Average spacing between Z-Bodies per cell in hiCMs from (H). (K) Number of α-actinin-2-positive puncta per cell in hiCMs from (A). (L) Average size of α-actinin-2-positive puncta in hiCMs from (A). (M) Representative images of α-actinin-2 and F-actin in control and Blebbistatin-treated hiCMs. (N) Myofibril orientation relative to the cell edge segment closest to the myofibril center, perpendicularly. N = 3 biological replicates, 668 control myofibrils, 385 50 µM Blebbistatin myofibrils, and 582 100 µM Blebbistatin myofibrils. Details of quantification can be found in Figure 2M–O.
α-actinin-2 quantification and organization in Blebbistatin-treated human induced pluripotent stem cell-derived cardiac myocytes (hiCMs) 12 hr post-plating.
(A) Myofibrils per cell in cells treated with DMSO or 50/100 µM Blebbistatin. N = 3 biological replicates, 95 control cells, 77 50 µM Blebbistatin cells, and 80 100 µM Blebbistatin cells. Myofibrils defined as having four or more Z-Lines in a row. More quantification details can be found in Figure 2—figure supplement 1. (B) Z-Lines per cell in hiCMs from (A). (C) Average myofibril persistence length per cell in hiCMs from (A): N = 3 biological replicates, 92 control cells, 69 50 µM Blebbistatin cells, and 65 100 µM Blebbistatin cells (only cells with myofibrils were quantified for C–E). (D) Average Z-Line length per cell in hiCMs from (C). (E) Average spacing between Z-Lines per cell in hiCMs from (C). (F) Muscle stress fibers (MSFs) per cell in hiCMs from (A). (G) Z-Bodies per cell in hiCMs from (A). (H) Average MSF persistence length per cell in hiCMs from (A): N = 3 biological replicates, 94 control cells, 77 50 µM Blebbistatin cells, and 80 100 µM Blebbistatin cells (only cells with MSFs were quantified for H–J). (I) Average Z-Body length per cell in hiCMs from (H). (J) Average spacing between Z-Bodies per cell in hiCMs from (H). (K) Number of α-actinin-2-positive puncta per cell in hiCMs from (A). (L) Average size of α-actinin-2-positive puncta in hiCMs from (A) .(M) Representative images of α-actinin-2 and F-actin in control and Blebbistatin-treated hiCMs. (N) Myofibril orientation relative to the cell edge segment closest to the myofibril center, perpendicularly. N = 3 biological replicates, 1560 control myofibrils, 1012 50 µM Blebbistatin myofibrils, and 777 100 µM Blebbistatin myofibrils. Details of quantification can be found in Figure 2M–O.
α-actinin-2 quantification and organization in Blebbistatin-treated human induced pluripotent stem cell-derived cardiac myocytes (hiCMs) 24 hr post-plating.
(A) Myofibrils per cell in cells treated with DMSO or 50/100 µM Blebbistatin. N = 4 biological replicates, 118 control cells, 108 50 µM Blebbistatin cells, and 93 100 µM Blebbistatin cells. Note that some of these subfigures were used also in Figure 3. Myofibrils defined as having four or more Z-Lines in a row. More quantification details can be found in Figure 2—figure supplement 1. (B) Z-Lines per cell in hiCMs from (A). (C) Average myofibril persistence length per cell in hiCMs from (A). (D) Average Z-Line length per cell in hiCMs from (B): N = 4 biological replicates, 104 control cells, 45 50 µM Blebbistatin cells, and 45 100 µM Blebbistatin cells (only cells with myofibrils were quantified for C–E). (E) Average spacing between Z-Lines per cell in hiCMs from (C). (F) Muscle stress fibers (MSFs) per cell in hiCMs from (A). (G) Z-Bodies per cell in hiCMs from (A). (H) Average MSF persistence length per cell in hiCMs from (A): N = 4 biological replicates, 118 control cells, 108 50 µM Blebbistatin cells, and 93 100 µM Blebbistatin cells (only cells with MSFs were quantified for H–J). (I) Average Z-Body length per cell in hiCMs from (H). (J) Average spacing between Z-Bodies per cell in hiCMs from (H). (K) Number of α-actinin-2-positive puncta per cell in hiCMs from (A). (L) Average size of α-actinin-2-positive puncta in hiCMs from (A).
sarcApp quantifies titin in hiCMs
Our next goal was to introduce additional quantitative modes of analysis to sarcApp that could be used concurrently with α-actinin-2-based quantification of Z-line assembly. Our focus was on other core components of the sarcomere which, like α-actinin-2, are thought to play an active role in assembly and can be stained and visualized using commercially available antibodies. Our group has shown previously that the protein titin can be used as a spatiotemporal indicator of myofibril maturation status in hiCMs (Taneja et al., 2020). Antibodies to the titin I-band localize to both MSFs and sarcomeres in hiCMs (Figure 4A). Titin forms ringlike structures around the Z-Bodies of MSFs that are closer to the apparent sarcomere transition point (Figure 4A). Figure 4B shows our current model for how titin is oriented around Z-bodies during earlier stages of sarcomere assembly, forming rings. Likely, due to both antibody localization to the titin I-Band region and the ability of titin to bind both actin and the α-actinin-2 N-terminus, titin is oriented with the N-terminus toward the center of the Z-Body and the C-terminus radially oriented outward (Figure 4C; Anderson and Granzier, 2012; Herzog, 2018). This is a topic of current research in our lab.
sarcApp uses titin binaries to identify myofibrils and precursor ring structures.
(A) Representative image of titin and α-actinin-2 in a control human induced pluripotent stem cell-derived cardiac myocyte (hiCM). Arrow: an α-actinin-2-positive Z-Body with titin localized in a ring around it. (B) Model of titin localization during sarcomere formation. (C) Titin doublets identified by sarcApp. Each line denotes a doublet with titin localized, and each color is a myofibril. (D) Quantification scheme for myofibrils and titin doublets. Details can be found in Figure 2—figure supplement 1 and the Methods. (E) Titin precursor rings identified by sarcApp (red). (F) Quantification scheme for titin precursor rings. Details can be found in Figure 2—figure supplement 1 and the Supplemental Methods.
yoU-Net-generated binaries of titin stains after training with annotated ground-truth binaries accurately recapitulate titin rings (MSFs) and titin doublets (sarcomeres) in hiCMs (Figure 4A, Figure 1—figure supplement 1D). We parameterized each titin-based structure into a quantitative scheme and have outfitted sarcApp with a modality for measuring titin. To determine the minimum length required for a potential titin doublet to be considered as part of a sarcomere, we co-stained hiCMs with both titin and α-actinin-2, and found that on average, titin doublets are 0.3 µm longer than the Z-Lines with which they associate. Thus, we set a lower boundary of titin doublet length at 1.7 µm (Z-lines: 1.4 µm). Closer inspection of titin images revealed that, in several cases, the titin I-Band signal ‘wraps’ around the radial tip of Z-Lines, reminiscent of titin rings around Z-Bodies.
sarcApp accurately distinguishes between titin doublets (Figure 4C) and titin rings (Figure 4E) and produces readouts for number of myofibrils doublets per cell, myofibril persistence length, doublet length, number of rings, ring diameter, ring aspect ratio, distance of doublets from the edge, and distance of rings from the edge. (Figure 4C–F). sarcApp is additionally equipped to incorporate a cell edge co-stain (e.g., actin or non-muscle myosin IIA or IIB) alongside titin to measure myofibril orientation and myofibril distance from edge since titin stains do not label the edge (similar to Figure 2M–O).
Blebbistatin affects myofibril orientation and doublet length, but not the periodicity of titin structures
We wished to use our sarcomere formation assay in the presence of Blebbistatin to quantify titin structure alignment using sarcApp. As before, hiCMs were replated in the presence of Blebbistatin for 24 hr, then fixed and stained for titin as well as phalloidin (F-actin) to aid in the visualization of MSFs. While F-actin in Blebbistatin-treated hiCMs is mostly punctate, titin rings and doublets can still be observed by eye around punctate actin with some degree of organization and periodicity (Figure 5A). sarcApp-dependent quantification of myofibrils using titin revealed no significant differences in the numbers of myofibrils in hiCMs replated in moderate (50 μM) or high (100 μM) Blebbistatin (Figure 5B), but found fewer and shorter titin doublets in high Blebbistatin (Figure 5C and D). sarcApp-based quantification of titin precursor rings within MSFs revealed no difference in absolute number of rings per cell with Blebbistatin treatment (Figure 5E) but rings that were detected were significantly more rounded as indicated by a decreased aspect ratio (Figure 5F). sarcApp also detected that myofibrils as quantified by titin were mostly parallel to the edge in control cells but became more randomly aligned relative to the edge in Blebbistatin (Figure 5G), consistent with sarcApp-dependent quantification of α-actinin-2. In summary, Blebbistatin treatment results in disarrayed and shorter titin doublets that are still capable of aligning into periodic myofibrillar structures.
Blebbistatin affects myofibril orientation and the morphology of titin structures.
(A) Representative images of titin and F-actin in human induced pluripotent stem cell-derived cardiac myocytes (hiCMs) treated with DMSO, 50 µM Blebbistatin, and 100 µM Blebbistatin. (B) Myofibrils per cell in hiCMs (N = 3 biological replicates, 107 DMSO cells, 95 50 µM Blebbistatin cells, and 84 100 µM Blebbistatin cells). (C) Doublets per cell in hiCMs from (B). (D) Doublet length per cell in hiCMs from (B) (N = 58 DMSO cells, 32 50 µM Blebbistatin cells, and 21 100 µM Blebbistatin cells). (E) Rings per cell in hiCMs from (B). (F) Average ring aspect ratio per cell in hiCMs from (B) (N = 105 DMSO control cells, 95 50 µM Blebbistatin cells, and 78 100 µM Blebbistatin cells). (G) Myofibril orientation N = 292 DMSO control myofibrils, 151 50 µM Blebbistatin myofibrils, and 69 100 µM Blebbistatin myofibrils. Full sarcApp outputs can be found in Figure 5—figure supplement 1.
Titin quantification and organization in Blebbistatin-treated human induced pluripotent stem cell-derived cardiac myocytes (hiCMs) 24 hr post-plating.
(A) Myofibrils per cell in cells treated with DMSO or 50/100 µM Blebbistatin. Myofibrils defined as having four or more doublets in a row. N = 3 biological replicates, 107 DMSO control cells, 95 50 µM Blebbistatin cells, and 84 100 µM Blebbistatin cells. (B) Titin doublets per cell. Only doublets within myofibrils are counted. (C) Myofibril persistence length average per cell in hiCMs from (A): N = 3 biological replicates, 58 control cells, 32 50 µM Blebbistatin cells, and 21 100 µM Blebbistatin cells (only cells with myofibrils were quantified for C–E and M–N). (D) Average doublet length per cell in hiCMs from (C). (E) Average spacing between doublets per cell in hiCMs from (C). (F) Number of titin precursor rings per cell in hiCMs from (A). (G) Average ring diameter per cell per cell in hiCMs from S6A: N = 3 biological replicates, 105 control cells, 95 50 µM Blebbistatin cells, and 78 100 µM Blebbistatin cells (only cells with rings were quantified for G–H and K–L). (H) Average ring aspect ratio per cell in hiCMs from (G). An aspect ratio of 1 is circular, and higher ratios are elongated. (I) Total number of titin-positive puncta per cell in hiCMs from (G). (J) Average size of all titin-positive puncta per cell in hiCMs from (A). (K) Average distance from the edge of the five closest titin rings, per cell, in cells from (H). (L) Average distance from the edge of all titin rings per cell, in cells from (H). (M) Average distance from the edge of the five closest titin doublets, per cell, in hiCMs from (C). (N) Average distance from the edge of all titin doublets within myofibrils per cell, in hiCMs from (C).
sarcApp quantifies myomesin, a component of the M-Line, in hiCMs
Having equipped sarcApp to quantify two structures associated with Z-Line assembly in hiCMs, our next goal was to introduce a modality for M-Line assembly. The M-Line lies at the midline of the stack of myosin II filaments in the sarcomere and demarcates the midpoint between two Z-Lines (Figure 6A). M-Lines in hiCMs can be stained using antibodies to the protein myomesin. Myomesin stains closely resemble α-actinin-2 Z-Line stains although myomesin is apparently absent from Z-Bodies (Figure 6B). Like α-actinin-2 and titin stains, yoU-Net can be used to predict myomesin binaries with high accuracy (Figure 6B, Figure 1—figure supplement 1E).
sarcApp uses myomesin binaries to identify myofibrils and M-Lines in human induced pluripotent stem cell-derived cardiac myocytes (hiCMs).
(A) Schematic showing myomesin localization at the M-Line. (B) Representative image of myomesin and F-actin in a hiCM. The myomesin binary was predicted using yoU-net as described in the Supplemental Methods. (C) Myofibrils and M-Lines identified by sarcApp. Each line denotes an M-Line, and each color represents a myofibril. (D) Quantification scheme for myofibrils and M-Lines. Details can be found in Figure 2—figure supplement 1. (E) Representative images of myomesin and F-actin in hiCMs treated with DMSO, 50 µM Blebbistatin, and 100 µM Blebbistatin. (F) Myofibrils per cell in hiCMs (N = 3 biological replicates; 112 DMSO cells, 90 50 µM Blebbistatin cells, and 89 100 µM Blebbistatin cells). (G) M-Lines per cell in hiCMs from (F). (H) Average M-Line length per cell in hiCMs from (F) (N = 97 DMSO control cells, 16 50 µM Blebbistatin cells, and 13 100 µM Blebbistatin cells). (I) Myofibril orientation (N = 752 DMSO control myofibrils, 49 50 µM Blebbistatin myofibrils, and 60 100 µM Blebbistatin myofibrils). Full sarcApp outputs can be found in Figure 6—figure supplement 1.
Myomesin quantification and organization in Blebbistatin-treated human induced pluripotent stem cell-derived cardiac myocyte (hiCMs) 24 hr post-plating.
(A) Myofibrils per cell in cells treated with DMSO or 50/100 µM Blebbistatin. Myofibrils defined as having three or more M-Lines in a row. N = 3 biological replicates, 112 DMSO control cells, 90 50 µM Blebbistatin cells, and 89 100 µM Blebbistatin cells. Quantification scheme details in Figure S2. (B) M-Lines per cell from (A). (C) Average myofibril persistence length per cell in hiCMs from (A): N = 3 biological replicates, 97 DMSO control cells, 16 50 µM Blebbistatin cells, and 13 100 µM Blebbistatin cells (only cells with myofibrils were quantified for C–E). (D) Average M-Line length per cell in hiCMs from (C). (E) Average spacing between M-Lines per cell in hiCMs from (C). (F) Number of myomesin-positive puncta per cell in hiCMs from (A). (G) Average size of all myomesin-positive puncta per cell in hiCMs from (A).
We equipped sarcApp to detect and measure myomesin M-Lines from yoU-Net-generated binaries (Figure 6C) based on a scheme that resembles the one designed for Z-Lines (Figure 6D). For myomesin, the M-Line must >1.4 µm and adjacent to at least two other M-lines to be considered part of a myofibril (i.e., at least three M-lines per myofibril, as it could be inferred that such a myofibril would have at least four Z-lines). sarcApp-defined readouts using myomesin include myofibrils per cell, M-Lines per cell, myofibril persistence length, and M-Line lengths. As with titin, sarcApp is additionally equipped to incorporate a cell edge co-stain (e.g., actin or non-muscle myosin IIA or IIB) alongside myomesin to measure myofibril orientation and myofibril distance from cell edge since myomesin antibodies do not stain the edge (see Figure 2M–O). To validate sarcApp as a tool to measure M-Lines, we stained myomesin in hiCMs exposed to Blebbistatin. Representative images in Figure 6E show that control hiCMs exhibit elongated M-Lines while Blebbistatin-treated M-Lines appear truncated and punctate-like. sarcApp quantification matched observations made by eye, detecting fewer myofibrils per cell (Figure 6F) and M-Lines per cell (Figure 6G) in Blebbistatin-treated hiCMs, as well as shortened M-Line length (Figure 6H). Co-staining with actin enabled sarcApp quantification of myofibril orientation, revealing that myofibril M-Lines are disorganized relative to the cell edge (Figure 6I, see Figure 2M–O) akin to the Blebbistatin-induced phenotype seen in Z-Lines (Figure 3H).
Neither α nor β cardiac myosin II alone is required for Z-Line assembly
The sarcomere A-band is the region of thick and thin filament overlap (Figure 7A). The thick filament contains stacks of muscle myosins that produce the forces of contraction (Henderson et al., 2017). Previous studies probing the role of muscle myosins in sarcomere assembly have produced conflicting results (Chopra et al., 2018; Yang et al., 2018). With sarcApp capable of measuring both M-Lines and Z-Lines, we were well-poised to ask if and/or how muscle myosins influence the specific assembly of both M-Lines and Z-Lines in hiCMs. Cardiac sarcomeres contain both α myosin II (MYH6) and β myosin II (MYH7) (Henderson et al., 2017; Chopra et al., 2018; Lyons et al., 1990); we exposed hiCMs to one of two unique sequences targeting either MYH- or MYH7 alongside a non-targeting siRNA (control) then replated, fixed, and stained hiCMs with antibodies to either α-actinin-2 (Z-Lines), titin (Z-Lines), or myomesin (M-Lines) (Figure 7B–C and H–I).
Knockdown of α or β cardiac myosin II reduces but does not eliminate sarcomeres.
(A) Schematic showing cardiac myosin localization in a sarcomere. (B) Representative western blot showing α cardiac myosin (MYH6) knockdown in hiCMs. (C) Representative images of α-actinin-2, titin, and myomesin in siControl human induced pluripotent stem cell-derived cardiac myocytes (hiCMs) and α cardiac myosin (MYH6) knockdown hiCMs. (D) Number of Z-Lines per cell in hiCMs in two independent groups of siControl (scramble)-treated hiCMs and two separate MYH6 siRNA-treated hiCMs (sequences 1 and 2). N = 3 biological replicates, 81 siCon cells and 78 siMYH6 (1) cells, and 88 siCon cells and 63 siMYH6 (2) cells. (E) Average Z-Line length per cell in hiCMs from (D). N = 68 siCon cells and 75 siMYH6 (1) cells, and 83 siCon cells and 62 siMYH6 (2) cells. (F) Average doublet length per cell in hiCMs in siCon (scramble)-treated hiCMs and two MYH6 siRNA sequences (1 and 2). 71 siCon cells, 49 siMYH6 (1) cells, and 50 siMYH6 (2) cells. (G) Average M-Line length per hiCMs. 85 siCon cells, 78 siMYH6 (1) cells, and 64 siMYH6 (2) cells. (H) Representative western blot showing β cardiac myosin (MYH7) knockdown in hiCMs. (I) Representative images of α-actinin-2, titin, and myomesin in β cardiac myosin (MYH7) knockdown hiCMs. (J) Number of Z-Lines per cell in hiCMs in two independent groups of siCon (scramble)-treated hiCMs and two separate MYH7 siRNA-treated hiCMs (pools 1 and 2). N = 86 siCon cells and 66 siMYH7 (1) cells, and 97 siCon cells and 62 siMYH7 (2) cells. (K) Average Z-Line length per cell in hiCMs from (J). 81 siCon cells and 63 siMYH7 (1) cells, and 81 siCon cells and 59 siMYH7 (2) cells. (L) Average doublet length per hiCM. N = 3 biological replicates, 94 siCon cells, 72 siMYH7 (1) cells, and 66 siMYH7 (2) cells. (M) Average M-Line length per cell in hiCMs. N = 87 siCon cells, 65 siMYH7 (1) cells, and 62 siMYH7 (2) cells. Full sarcApp outputs and western blots can be found in Figure 7—figure supplements 1–7.
Original 16-bit image file for anti-MYH6 western blot.
See Figure 7-figure supplement 7A and Figure 7B.
Original 16-bit image file for anti-tubulin western blot.
See Figure 7-figure supplement 7A and Figure 7B.
Original 16-bit image file for anti-MYH7 western blot.
See Figure 7-figure supplement 7B and Figure 7H.
Original 16-bit image file for anti-tubulin western blot.
See Figure 7-figure supplement 7B and Figure 7H.
α-actinin-2 quantification and organization in MYH6 knockdown human induced pluripotent stem cell-derived cardiac myocytes (hiCMs).
(A) Myofibrils per cell in two groups of siCon (scramble)-treated hiCMs and two separate MYH6 siRNA-treated hiCMs (sequences 1 and 2). N = 3 biological replicates, 81 siCon cells and 78 siMYH6 (1) cells, and 88 siCon cells and 63 siMYH6 (2) cells. Myofibrils defined as having four or more Z-Lines in a row. Quantification details can be found in Figure 2—figure supplement 1. (B) Z-Lines per cell in hiCMs from (A). (C) Average myofibril persistence length per cell in hiCMs from (B): N = 68 siCon cells and 75 siMYH6 (1) cells, and 83 siCon cells and 62 siMYH6 (2) cells (only cells with myofibrils were quantified for C–E). (D) Average Z-Line length per cell in hiCMs from (C). (E) Average spacing between Z-Lines per cell in hiCMs from (C). (F) Muscle stress fibers (MSFs) per cell in cells from (A). (G) Z-Bodies per cell in hiCMs from (A). (H) Average MSF persistence length per cell in hiCMs from (A): N = 3 biological replicates, X7 siCon cells, X8 siMYH6 (1) cells, and X9 siMYH6 (2) cells (only cells with MSFs were quantified for H–J). (I) Average Z-Body length per cell in hiCMs from (H). (J) Average spacing between Z-Bodies per cell in hiCMs from (H). (K) Number of α-actinin-2-positive puncta per cell in hiCMs from (A). (L) Average size of α-actinin-2-positive puncta per cell in hiCMs from (A). (M) Myofibril orientation relative to the cell edge segment closest to the myofibril center, perpendicularly. N = 3 biological replicates: 898 siCon myofibrils and 1173 siMYH6 (1) myofibrils, and 964 siCon myofibrils and 974 siMYH6 (2) myofibrils. Details of quantification can be found in Figure 2M–O. N = 3 biological replicates.
Titin quantification and organization in MYH6 knockdown human induced pluripotent stem cell-derived cardiac myocytes (hiCMs).
(A) Myofibrils per cell in siCon (scramble)-treated hiCMs and two separate MYH6 siRNA-treated hiCMs (sequences 1 and 2). Myofibrils were defined as having four or more doublets in a row. N = 3 biological replicates, 91 siCon cells, 73 siMYH6 (1) cells, and 66 siMYH6 (2) cells. (B) Titin doublets per cell from Figure S9A. (C) Myofibril persistence length average per cell in hiCMs from (A): N = 3 biological replicates, 71 siCon cells, 49 siMYH6 (1) cells, and 50 siMYH6 (2) cells (only cells with myofibrils were quantified for C–E and M–N). (D) Average doublet length per cell in hiCMs from (C). (E) Average spacing between doublets per cell in hiCMs from (C). (F) Number of titin precursor rings per cell in hiCMs from (A). (G) Average ring diameter per cell in hiCMs from (A). N = 3 biological replicates, 78 siCon cells, 67 siMYH6 (1) cells, 60 siMYH6 (2) cells (only cells with rings were quantified for G–H and K–L). (H) Average ring aspect ratio per cell in hiCMs from (G). An aspect ratio of 1 is circular, and higher ratios are elongated. (I) Total number of titin-positive puncta per cell in hiCMs from (A). (J) Average size of all titin-positive puncta per cell in hiCMs from (A). (K) Average distance from the edge of the five closest titin rings, per cell, in hiCMs from (G). (L) Average distance from the edge of all titin rings per cell in hiCMs from (G). (M) Average distance from the edge of the five closest titin doublets, per cell, in hiCMs from (C). (N) Average distance from the edge of all titin doublets within myofibrils per cell in hiCMs from (C). (O) Myofibril orientation relative to the cell edge segment closest to the myofibril center, perpendicularly in hiCMs from (B). N = 3 biological replicates, 449 siCon myofibrils, 440 siMYH6 (1) myofibrils, and 442 siMYH6 (2) myofibrils. Details of quantification can be found in Figure 2M–O.
Myomesin quantification and organization in MYH6 knockdown cells.
(A) Myofibrils per cell in siCon (scramble)-treated human induced pluripotent stem cell-derived cardiac myocytes (hiCMs) and two separate MYH6 siRNA-treated hiCMs (sequences 1 and 2). Myofibrils defined as having three or more M-Lines in a row. N = 3 biological replicates, 100 siCon cells, 80 siMYH6 (1) cells, and 71 siMYH6 (2) cells. Quantification scheme details in Figure 2—figure supplement 1. (B) M-Lines per cell from (A). (C) Average myofibril persistence length per cell in hiCMs from (A): N = 3 biological replicates, 85 siCon cells, 78 siMYH6 (1) cells, and 64 siMYH6 (2) cells (only cells with myofibrils were quantified for C–E). (D) Average M-Line length per cell in hiCMs from (C). (E) Average spacing between M-Lines per cell in hiCMs from (C). (F) Number of myomesin-positive puncta per cell in hiCMs from (A). (G) Average size of all myomesin-positive puncta per cell in hiCMs from (A). (H) Myofibril orientation relative to the cell edge segment closest to the myofibril center, perpendicularly. N = 3 biological replicates, 685 siCon myofibrils, 979 siMYH6 (1) myofibrils, and 743 siMYH6 (2) myofibrils. Details of quantification can be found in Figure 2M–O.
α-actinin-2 quantification and organization in MYH7 knockdown human induced pluripotent stem cell-derived cardiac myocytes (hiCMs).
(A) Myofibrils per cell in two groups of siCon (scramble)-treated hiCMs and two separate MYH7 siRNA-treated hiCMs (pools 1 and 2). N = 3 biological replicates, 86 siCon cells and 66 siMYH7 (1) cells, and 97 siCon cells and 62 siMYH7 (2) cells. Myofibrils were defined as having four or more Z-Lines in a row. Details can be found in Figure 2—figure supplement 1. (B) Z-Lines per cell in hiCMs from (A). (C) Average myofibril persistence length per cell in hiCMs from (A): N = 3 biological replicates, 81 siCon cells and 62 siMYH7 (1) cells, and 81 siCon cells and 59 siMYH7 (2) cells (only cells with myofibrils were quantified for C–E). (D) Average Z-Line length per cell in hiCMs from (C). (E) Average spacing between Z-Lines per cell in hiCMs from (C). (F) Muscle stress fibers (MSFs) per cell in cells from (A). (G) Z-Bodies per cell in hiCMs from (A). (H) Average MSF persistence length per cell in hiCMs from (A): N = 3 biological replicates, 86 siCon cells and 66 siMYH7 (1) cells, and 97 siCon cells and 62 siMYH7 (2) cells (only cells with MSFs were quantified for H–J). (I) Average Z-Body length per cell in hiCMs from (H). (J) Average spacing between Z-Bodies per cell in hiCMs from (H). (K) Number of α-actinin-2-positive puncta per cell in hiCMs from (A). (L) Average size of α-actinin-2-positive puncta per cell in hiCMs from (A). (M) Myofibril orientation relative to the cell edge segment closest to the myofibril center, perpendicularly. N = 3 biological replicates, 878 siCon myofibrils and 1126 siMYH7 (1) myofibrils, and 622 siCon myofibrils and 1324 siMYH7 (2) myofibrils. Details of quantification can be found in Figure 2M–O.
Titin quantification and organization in MYH7 knockdown human induced pluripotent stem cell-derived cardiac myocytes (hiCMs).
(A) Myofibrils per cell in siCon (scramble)-treated hiCMs and two separate MYH7 siRNA-treated hiCMs (pools 1 and 2). Myofibrils defined as having four or more doublets in a row. N = 3 biological replicates, 94 siCon cells, 72 siMYH7 (1) cells, and 66 siMYH7 (2) cells. (B) Titin doublets per cell from (A). (C) Myofibril persistence length average per cell in hiCMs from (A): N = 3 biological replicates, 56 siCon cells, 56 siMYH7 (1) cells, and 52 siMYH7 (2) cells (only cells with myofibrils were quantified for C–E and M–N). (D) Average doublet length per cell in hiCMs from (C). (E) Average spacing between doublets per cell in hiCMs from (C). (F) Number of titin precursor rings per cell in hiCMs from (A). (G) Average ring diameter per cell in hiCMs from (A): N = 3 biological replicates, 90 siCon cells, 58 siMYH7 (1) cells, and 64 siMYH7 (2) cells (only cells with rings were quantified for G–H and J-K). (H) Average ring aspect ratio per cell in hiCMs from (G). An aspect ratio of 1 is circular, and higher ratios are elongated. (I) Total number of titin-positive puncta per cell in hiCMs from (A). (J) Average size of all titin-positive puncta per cell in hiCMs from (A). (K) Average distance from the edge of the five closest titin rings, per cell, in hiCMs from (G). (L) Average distance from the edge of all titin rings per cell in hiCMs from (G). (M) Average distance from the edge of the five closest titin doublets, per cell, in hiCMs from (C). (N) Average distance from the edge of all titin doublets within myofibrils per cell in hiCMs from (C). (O) Myofibril orientation relative to the cell edge segment closest to the myofibril center, perpendicularly in hiCMs from (B). N = 3 biological replicates, 278 siCon myofibrils, 318 siMYH7 (1) myofibrils, and 458 siMYH7 (2) myofibrils. Details of quantification can be found in Figure 2M–O.
Myomesin quantification and organization in MYH7 knockdown cells.
(A) Myofibrils per cell in siCon (scramble)-treated human induced pluripotent stem cell-derived cardiac myocytes (hiCMs) and two separate MYH7 siRNA-treated hiCMs (pools 1 and 2). Myofibrils defined as having three or more M-Lines in a row. N = 3 biological replicates, 104 siCon cells, 75 siMYH7 (1) cells, and 70 siMYH7 (2) cells. Quantification scheme details in Figure 2—figure supplement 1. (B) M-Lines per cell in hiCMs from (A). (C) Average myofibril persistence length per cell in hiCMs from (A): N = 3 biological replicates, 87 siCon cells, 65 siMYH7 (1) cells, and 62 siMYH7 (2) cells (only cells with myofibrils were quantified for Figure S13C–E). (D) Average M-Line length per cell in hiCMs from (C). (E) Average spacing between M-Lines per cell in hiCMs from (C). (F) Number of myomesin-positive puncta per cell in hiCMs from (A). (G) Average size of all myomesin-positive puncta per cell in hiCMs from (A). (H) Myofibril orientation relative to the cell edge segment closest to the myofibril center, perpendicularly. N = 3 biological replicates, 793 siCon myofibrils, 775 siMYH7 (1) myofibrils, and 778 siMYH7 (2) myofibrils. Details of quantification can be found in Figure 2M–O.
MYH6 and MYH7 knockdown western blots.
(A) Full image for the western blots shown in in Figure 7B. Individual lanes are denoted by numbers. Protein knockdown using siRNA targeted to MYH6. (C) Full image for the western blots shown in in Figure 7H. Individual lanes are denoted by numbers. (D) Protein knockdown using siRNA targeted to MYH7. N = 2 biological replicates. Anti-MYH6 and Anti-MYH7 images displayed at 0–50,000 gray levels. Anti-tubulin images displayed at 0–40,000. Full 16-bit images can be found in Figure 7—source data 1–4.
After 24 hr, we observed by eye that both the Z-Lines and M-Lines of hiCMs depleted of α myosin II (MYH6) appeared to largely resemble those of controls (Figure 7C). sarcApp-dependent quantification of each stain determined that α myosin II knockdown did not alter length of α-actinin-2 Z-lines, titin doublets, or M-lines, though one of the two siRNAs did result in increased numbers of Z-lines (Figure 7D–G). As with α myosin II, stains of β myosin II-depleted hiCMs (MYH7) also appeared to resemble those of controls by eye (Figure 7H and I); however, sarcApp-based quantification revealed that hiCMs depleted of β myosin II assembled more abundant, but shorter Z-lines than controls (Figure 7J and K). Further, while sarcApp detected no change in titin doublet lengths (Figure 7L), M-lines were detected as being significantly shorter than controls following β myosin II knockdown (Figure 7M). Thus, we conclude that neither α nor β myosin II is strictly required for Z- or M-line assembly in hiCMs but that β myosin II likely influences Z-line concatenation. This is not entirely surprising, as Geach and colleagues showed similar results in Xenopus embryos (Geach et al., 2015).
Myomesin is required for A-band but not Z-Line assembly
Given our data suggesting that Z-Line and M-Line assembly could be uncoupled (Figure 7), we next asked if directly disrupting M-Line assembly impacts Z-Line assembly. We disrupted M-Line assembly by exposing hiCMs to one of two types of myomesin-targeting siRNAs (gene name: MYOM) alongside a non-targeting siRNA (control) then replated, fixed, and stained hiCMs with antibodies to either α-actinin-2 or titin (Figure 8A and B).
Myomesin knockdown alters titin and cardiac myosin II localization, but not α-actinin-2.
(A) Representative images of sarcomeric proteins α-actinin-2 and titin in myomesin (MYOM) knockdown human induced pluripotent stem cell-derived cardiac myocytes (hiCMs). (B) Representative western blot and quantification showing MYOM knockdown in hiCMs, N = 3 biological replicates. (C) Number of Z-Lines per cell in siCon (scramble)-treated hiCMs and two separate MYOM siRNA-treated hiCMs (sequences 1 and 2). N = 3 biological replicates, 132 siCon cells, 105 siMYOM (1) cells, and 103 siMYOM (2) cells. (D) Average Z-Line length per cell in hiCMs from (C): N = 4 biological replicates, 117 siCon cells, 104 siMYOM (1) cells, and 92 siMYOM (2) cells (only cells with myofibrils were quantified for D). (E) Titin doublets per cell in hiCMs using experimental treatments in (C). N = 3 biological replicates, 117 siCon cells, 90 siMYOM (1) cells, and 100 siMYOM (2) cells. (F) Average doublet length per cell in hiCMs from (E): N = 3 biological replicates, 74 siCon cells, 68 siMYOM (1) cells, and 70 siMYOM (2) cells (only cells with myofibrils were quantified for F). (G) Rings per cell in hiCMs from (E). (H) Representative images of MYH7 and F-actin in siCon, siMYOM (1), and siMYOM (2) hiCMs. (I) Length of β cardiac myosin stacks in siCon, siMYOM (1), and siMYOM (2) hiCMs. Full sarApp outputs and western blot can be found in Figure 8—figure supplements 1–3.
Original 16-bit image file for anti-MYH6 western blot.
See Figure 8—figure supplement 3A and Figure 8B.
Original 16-bit image file for anti-tubulin western blot.
See Figure 8—figure supplement 3A and Figure 8B.
α-actinin-2 quantification and organization in myomesin knockdown human induced pluripotent stem cell-derived cardiac myocytes (hiCMs).
(A) Myofibrils per cell in siCon (scramble)-treated hiCMs and two separate myomesin siRNA-treated hiCMs (sequences 1 and 2). Myofibrils defined as having four or more Z-Lines in a row. N = 4 biological replicates, 132 siCon cells, 105 siMYOM (1) cells, and 103 siMYOM (2) cells. More quantification details can be found in Figure 2—figure supplement 1. (B) Z-Lines per cell in hiCMs from (A). (C) Average myofibril persistence length per cell in hiCMs from (A): N = 4 biological replicates, 117 siCon cells, 104 siMYOM (1) cells, and 92 siMYOM (2) cells (only cells with myofibrils were quantified for C–E). (D) Average Z-Line length per cell in hiCMs from (C). (E) Average spacing between Z-Lines per cell in hiCMs from (C). (F) Muscle stress fibers (MSFs) per cell in cells from (A). (G) Z-Bodies per cell in hiCMs from (A). (H) Average MSF persistence length per cell in hiCMs from Figure S14A: N = 4 biological replicates, 131 siCon cells, 105 siMYOM (1) cells, and 103 siMYOM (2) cells (only cells with MSFs were quantified for H–J). (I) Average Z-Body length per cell in hiCMs from (H). (J) Average spacing between Z-Bodies per cell in hiCMs from (H). (K) Number of α-actinin-2-positive puncta per cell in hiCMs from (A). (L) Average size of α-actinin-2-positive puncta per cell in hiCMs from (A). (M) Myofibril orientation relative to the cell edge segment closest to the myofibril center, perpendicularly. N = 4 biological replicates, 862 siCon myofibrils, 1305 siMYOM (1) myofibrils, and 1086 siMYOM (2) myofibrils. Details of quantification can be found in Figure 2M–O.
Titin quantification and organization in myomesin knockdown human induced pluripotent stem cell-derived cardiac myocytes (hiCMs).
(A) Myofibrils per cell in siCon (scramble)-treated hiCMs and two separate MYOM siRNA-treated hiCMs (sequences 1 and 2). Myofibrils defined as having four or more doublets in a row. N = 3 biological replicates, 117 siCon cells, 90 siMYOM (1) cells, and 100 siMYOM (2) cells. More quantification details can be found in Figure 2—figure supplement 1. (B) Titin doublets per cell in hiCMs from (A). (C) Myofibril persistence length average per cell in hiCMs from (A): N = 3 biological replicates, 74 siCon cells, 68 siMYOM (1) cells, and 70 siMYOM (2) cells (only cells with myofibrils were quantified for C–E and M–N). (D) Average doublet length per cell in hiCMs from (C). (E) Average spacing between doublets per cell in hiCMs from (C). (F) Number of titin precursor rings per cell in hiCMs from (A). (G) Average ring diameter per cell in hiCMs from (A): N = 3 biological replicates, 106 siCon cells, 71 siMYOM (1) cells, and 89 siMYOM (2) cells. (Only cells with rings were quantified for G-H and K, L). (H) Average ring aspect ratio per cell in hiCMs from (G). An aspect ratio of 1 is circular, and higher ratios are elongated. (I) Total number of titin-positive puncta per cell in hiCMs from (A). (J) Average size of all titin-positive puncta per cell in hiCMs from (A). (K) Average distance from the edge of the five closest titin rings, per cell, in hiCMs from (G). (L) Average distance from the edge of all titin rings per cell in hiCMs from (G). (M) Average distance from the edge of the five closest titin doublets, per cell, in hiCMs from (B). (N) Average distance from the edge of all titin doublets within myofibrils per cell in hiCMs from (B). (O) Myofibril orientation relative to the cell edge segment closest to the myofibril center, perpendicularly in hiCMs from (B). N = 3 biological replicates, 361 siCon myofibrils, 561 siMYOM (1) myofibrils, and 667 siMYOM (2) myofibrils. Details of quantification can be found in Figure 2M–O.
MYOM knockdown western blots.
(A) Full image for the western blots shown in in Figure 8B. Individual lanes are denoted by numbers. Protein knockdown using siRNA targeted to MYOM. Individual lanes are denoted by numbers. (B) Protein knockdown using siRNA targeted to MYH7. N = 3 biological replicates. Anti-MYOM image displayed at 0–50,000 gray levels. Anti-tubulin image displayed at 0–40,000. Full 16 bit images can be found in Figure 8—source data 1 and 2.
After 24 hr, we observed by eye that both the α-actinin-2 and titin stains of myomesin-depleted hiCMs resemble those of controls (Figure 8A). sarcApp-dependent quantification of α-actinin-2 detected no difference in Z-Lines per cell in myomesin-depleted hiCMs compared to controls (Figure 8C), suggesting Z-Lines assemble independently of myomesin; however, sarcApp did detect a decrease in Z-Line length for one myomesin-targeting siRNA (Figure 8D, Figure 8—figure supplement 1). Meanwhile sarcApp-dependent quantification of titin detected no significant change in the number of titin doublets or doublet lengths (Figure 8E and F). Interestingly, upon myomesin knockdown, the number of titin precursor rings significantly increase, warranting future investigation (Figure 8G).
We next wanted to assess the influence of myomesin knockdown on M-Line/A-band assembly. Because we were unable to visualize M-Lines in myomesin-depleted hiCMs (data not shown), we stained the A-band using antibodies to the β myosin II motor domain, observing that A-bands in myomesin-depleted hiCMs appear to be less organized (Figure 8H) with shorter β myosin II stacks (Figure 8I). These data suggest that myomesin regulates A-band/M-Line assembly and myofibril maturation, but not Z-Line assembly. Altogether, these data in tandem with data from Figure 7 suggest that the assembly of the sarcomere A-band/M-Line and Z-Line are decoupled and/or regulated by mechanisms that are at least partially exclusive to one another.
Discussion
Here, we introduce sarcApp and yoU-Net, two open-source software packages to binarize microscope images and quantify sarcomere components. We first demonstrate deep learning-based image binarization of three distinct sarcomeric proteins – α-actinin-2, titin, and myomesin – using models trained by a framework we developed called yoU-Net, based off the ubiquitously used U-Net (Ronneberger and Brox, 2015). Each trained model is provided and will function as demonstrated herein based on the use of the antibodies as stated in the ‘Methods.’ yoU-Net is designed and equipped to be functionally versatile, which we hope other researchers can harness to generate additional trained models of proteins not studied here.
The high fidelity of yoU-Net-generated binaries allowed us to quantify several geometric features of the sarcomere in real space as opposed to frequency space. Automatic quantification of sarcomere/myofibril structure with sarcApp offers multiple advantages over traditional, manual quantification including speed, objectivity, and consistency. sarcApp is capable of quantifying 33 measurements per cell and 24 measurements per myofibril at a rate that is orders of magnitude faster than manual quantification, which besides being slower also requires complete attention from the user. sarcApp also removes the potential for user bias/error from quantification and can detect quantitative differences in cells that are not readily apparent by eye (Figures 7 and 8) – we expect this will drive discovery of novel mechanisms in cells that would have remained otherwise undetectable. sarcApp as an automatic tool must operate within a rigid quantitative framework and functions optimally when structures o -interest have shapes that are consistent and definable by simple geometric measurements; however, to increase usability we have enabled users to self-define parameters using an easy-to-navigate GUI.
Other than basic structural readouts for sarcomere/myofibril precursors (Z-Bodies, titin rings, etc.) and sarcomeres/myofibrils (Z-Lines, titin doublets, M-Lines, etc.), sarcApp is also equipped to proxy sarcomere/myofibril organization. Myofibril organization correlates with function in vivo and poor organization (disarray) is often cited as causative for myocardial disease (Eschenhagen et al., 2015; Harvey and Leinwand, 2011; Olsson et al., 2004). sarcApp reports organization using a novel metric reported here known as myofibril orientation (Figure 2M–O). Orientation is reported relative to the cell membrane by comparing the angle of the myofibril long axis to the nearest cell edge; such a metric was chosen because we routinely observe hiCMs assemble myofibrils with a long axis that is approximately parallel (i.e., 0°) to the cell edge in vitro and also because tissue sections of myocytes in vivo show myofibrillar arrays parallel to the long axis of the membrane (Fenix et al., 2018; Taneja et al., 2020). We generated this measure of orientation to be more transparent and informative than what is currently available in the hopes of reaching a broader audience (Chopra et al., 2018; Hinson et al., 2015).
Previous work from our lab and others has included as many cells as reasonably possible in analyses (Fenix et al., 2018; Taneja et al., 2020), requiring technical expertise in (at minimum) plating cells, sample preparation, microscopy, and image analysis. While microscopy has historically been the rate-limiting step in such analyses, modern microscopes can now rapidly image hundreds to thousands of cells at high resolution. The true bottleneck towards understanding sarcomere assembly now lies for our lab in image analysis/quantification. We summed the quantified cell totals of two recent landmark papers in the field and found the reported totals to be 385 (Chopra et al., 2018) and 785 (Fenix et al., 2018). sarcApp has allowed us to present in this article multidimensional outputs from 3452 single cells with outputs that we feel represent the data clearly at both basic glance and upon close inspection. Much like yoU-net, we have encoded sarcApp into a GUI equipped with a user manual that details the steps one needs to use sarcApp, irrespective of coding expertise.
We validated sarcApp using the pan-myosin II inhibitor Blebbistatin (Limouze et al., 2004). Blebbistatin exposure results in predictable, dose-dependent inhibition of sarcomere assembly (Chopra et al., 2018; Liu et al., 2016). sarcApp-generated measurements of α-actinin-2 in the presence or absence of Blebbistatin were consistent with observations made by previous groups (Chopra et al., 2018; Liu et al., 2016; Figure 3). Meanwhile, several other outputs related to titin and myomesin first reported here by sarcApp suggest that periodic, pseudomyofibrillar structures characteristic to both the Z-Line and M-Line can at least partially self-assemble in the absence of myosin II motor-based contractility (Figures 5 and 6). These data indicate other mechanisms beyond myosin II-based motor-based contractility may independently contribute to the assembly and ordered arrangement of myofibrils in hiCMs. Specifically with titin, we also observe the phenomenon that Blebbistatin disrupts the higher-order organization of titin-based structures with respect to the cell membrane but does not affect titin periodicity within the structure. These data suggest that unique mechanistic tiers of organization exist within the cell at the sub- and super-structural level that become uncoupled upon relieving the cell of myosin II-dependent contractility.
We find that neither α nor β cardiac myosin II knockdown had any measurable impact on the assembly of Z-Lines (Figure 7). Attempts to determine if Z-Lines assemble in the simultaneous absence of both myosins were unsuccessful in our hands. Such an experiment, in addition to the Blebbistatin experiments reported herein and elsewhere (Chopra et al., 2018; Liu et al., 2016), could resolve whether myosin II-dependent contractility alone is sufficient to explain the role of myosin II in Z-Line assembly. Meanwhile, β cardiac myosin II knockdown did result in hiCMs with shorter M-Lines, indicating that either contractility or scaffolding by β, but not α myosin II, is required for M-Line maturation (Figure 7G and M). Knockdown of myomesin similarly did not impact Z-Line assembly but did result in shortening of the thick filament A-band (Figure 8H and I). Our ability to perturb the M-Line/A-band but not the Z-Line suggests that Z- and M-Line assembly are regulated at least in part by distinct mechanistic axes, as previously hypothesized (Rudy et al., 2001; Schafer et al., 1995; Beall et al., 1989). It should be noted here that solely using Z-Lines as a proxy for sarcomere status would have led us to errantly conclude that β cardiac myosin II and myomesin both play no or at least minor roles in sarcomere assembly. Future studies examining assembly should therefore consider Z-Line assembly to be distinct from, or at least not wholly representative of, assembly of other sarcomere components.
Here we used exclusively fixed-cell immunofluorescence for high-throughput quantification of sarcomere assembly. We eliminated as much bias as possible by selecting cells for quantification solely using a nuclear DAPI stain and are therefore confident that data presented herein fully captures the true biological variability present within whole hiCM cultures. A future goal of the lab is to further adapt sarcApp for live cell quantification. Tracking specific structures across time would be a necessity for a ‘live’ version of sarcApp, a hurdle the lab is currently working to overcome. We also designed this initial version of sarcApp to quantify in 2D rather than in 3D space. As we have published previously, sarcomeres/myofibrils assemble on the dorsal (top) surface of an hiCM plated on glass for up to 48 hr after plating, and we consider them within that timeframe to exist largely within a two-dimensional plane (Fenix et al., 2018; Taneja et al., 2020). Breaking into three dimensions will require more complex mathematics and produce more convoluted outputs, which could limit adoption by other researchers, but is a long-term goal of the lab nonetheless.
In summary, sarcApp and yoU-Net provide frameworks for automatic binarization, segmentation, and quantitative analysis of sarcomeric proteins in striated muscle cells. We anticipate these software packages will be useful for lower-throughput single-cell analyses of sarcomere structure and/or assembly as well as for high-throughput screens to analyze several hundreds to thousands of cells at a time. In this way, we hope to facilitate the entry of more researchers into the field of cardiac cell biology and continue to introduce novel quantification metrics and additional sarcomere proteins in the future through continued maintenance and management of sarcApp as an open-source community project.
Methods
SarcApp and yoU-net can be found at the following address:
https://github.com/abbieneininger/sarcApp; (copy archived at Abbieneininger, 2023).
Reagent type (species) or resource
Designation
Source or reference
Identifiers
Additional information
Antibody
Anti-alpha actinin 2 (mouse monoclonal)
Sigma
A7811
IF, 1:200
Antibody
Anti-Titin (mouse monoclonal)
DHSB
9D10
IF, 1:2
Antibody
Mouse anti-Myomesin
DHSB
MYOM
IF, 1:2
Antibody
Mouse anti-MYH
DHSB
A4.591
IF, 1:2
Antibody
Rabbit anti-MYH6
ProteinTech
22281-1-AP
WB, 1:500
Antibody
Rabbit anti-MYH7
ProteinTech
22280-1-AP
WB, 1:500
Antibody
Goat anti-mouse 488
Life Technologies
A11001
IF, 1:100
Antibody
Goat anti-rabbit 488
Life Technologies
A11034
IF, 1:100
Antibody
Goat anti-mouse 568
Life Technologies
A11004
IF, 1:100
Antibody
Goat anti-rabbit 568
Life Technologies
A11036
IF, 1:100
Antibody
Goat anti-mouse 647
Life Technologies
A32728
IF, 1:100
Antibody
Goat anti-rabbit 647
Life Technologies
A32733
IF, 1:100
Biological sample (Bos taurus)
Bovine serum albumin
RPI
A30075-100
Chemical compound, drug
Phalloidin Alexa Fluor 488
Invitrogen
A12379
Chemical compound, drug
Phalloidin Alexa Fluor 568
Invitrogen
A12380
Chemical compound, drug
Phalloidin Alexa Fluor 647
Invitrogen
A22287
Chemical compound, drug
PBS, 10×, Ca2+/Mg2+ free
Life Technologies
70011-044
Chemical compound, drug
PBS, 10×, with Ca2+/Mg2+
Corning
46-013CM
Chemical compound, drug
PFA, 16%
Electron Microscopy Sciences
15710
Chemical compound, drug
0.5% Trypsin
Life Technologies
15400-054
Chemical compound, drug
0.1% Gelatin
Sigma
ES-006-B
Chemical compound, drug
Dimethyl sulfoxide
Sigma
276855
Chemical compound, drug
Vectashield with DAPI
Vector
H-1200
Chemical compound, drug
Blebbistatin
Sigma
B0560
Chemical compound, drug
Fibronectin
Corning
354008
Chemical compound, drug
Lipofectamine RNAiMAX
Thermo Fisher
LMRNA015
Chemical compound, drug
TBS, 10×
Corning
46-012CM
Chemical compound, drug
Tween 20
Sigma
P7949
Cell line (Homo sapiens)
iCell cardiomyocytes^2 kit
Fujifilm International
CMC-100-012-000.5
Sequence-based reagent
SMART Pool siRNA against human MYH7 (13–16)
Horizon Discovery
A-011086
Sequence-based reagent
SMART Pool siRNA against human MYH7 (13, 14)
Horizon Discovery
A-011086-13, A-011086-14
Sequence-based reagent
siRNA against human MYH6 3’ UTR (13)
Horizon Discovery
A-012645-13-0005
Sequence-based reagent
siRNA against human MYH6 CDS (14)
Horizon Discovery
A-012645-14-0005
Sequence-based reagent
siRNA against human myomesin 3’UTR (13)
Horizon Discovery
A-006342-13-0005
Sequence-based reagent
siRNA against human myomesin CDS (16)
Horizon Discovery
A-006342-16-0005
Software, algorithm
FIJI
NIH
Software, algorithm
sarcApp
This study
Software, algorithm
yoU-Net
This study
The supplemental user manual and sarcApp software
Details on sarcApp and yoU-Net (Neininger-Castro, 2023) download and usage documentation are given in the supplement and at https://github.com/abbieneininger/sarcApp; (copy archived at Abbieneininger, 2023). Image data is available upon request.
Cell culture and authentication
Human iPSC-derived cardiac myocytes (CMM-100-012-000.5, Cellular Dynamics, Madison, WI) were cultured as per the manufacturer’s instructions in proprietary manufacturer-provided cardiac myocyte maintenance medium in polystyrene 96-well cell culture plates. Cells were maintained at 37°C and 5% CO2. For replating hiCMs onto glass substrates, cells were washed two times with 100 µL 1× PBS with no Ca2+/Mg2+ (PBS*, 70011044, Gibco, Grand Island, NY). PBS* was completely removed from hiCMs and 40 µL 0.1% Trypsin-EDTA with no phenol red (15400054, Gibco) was added to hiCMs and incubated at 37°C for 2 min. Following incubation, the cells were washed three times with trypsin, the plate rotated 180°, and washed another three times. Trypsinization was then quenched by adding 120 µL of culture media and total cell mixture was pipetted into a 1.5 mL Eppendorf tube. Cells were centrifuged at 200 × g for 3 min, and the supernatant was aspirated. The cell pellet was resuspended in 200 µL of culture media and plated on 35 mm dishes with a 10 mm glass bottom (D35-10-1.5-N; CellVis, Sunnydale, CA) pre-coated with 10 µg/mL fibronectin (354008, Corning) for 1 hr at 37°C.
Antibodies
Alexa Fluor 488-phalloidin (A12379), Alexa Fluor 568-phalloidin (A12380), and Alexa Fluor 647-phalloidin (A22287) were purchased from Invitrogen. Alexa Fluor 488-goat anti-mouse (A11029), Alexa Fluor 488-goat anti-rabbit (A11034), Alexa Fluor 568-goat-anti-rabbit (A11011), Alexa Fluor 568-goat anti-mouse (A11004), Alexa Fluor 647-goat-anti-mouse (A32728), and Alexa Fluor 647-goat-anti-rabbit (A32733) (1:100) antibodies were purchased from Life Technologies (Grand Island, NY).
The Titin (9D10) antibody, Myomesin (MYOM) antibody, and MYH7 (A4.591) (all 1:2) antibody were purchased from the Developmental Studies Hybridoma Bank (University of Iowa). Mouse anti-α-actinin-2 (1:200, A7811) was purchased from Sigma-Aldrich. Rabbit anti-MYH6 and MYH7 for western blotting (1:500) were purchased from ProteinTech (22281-1-AP, 22280-1-AP).
Chemicals
Blebbistatin was purchased from Sigma (B0560) and reconstituted to 10 mM in DMSO (Sigma 276855).
Fixation and Immunostaining
Cells were fixed with 4% paraformaldehyde (PFA) in PBS at room temperature for 20 min and then permeabilized for 5 min with 1% Triton X-100/4% PFA in PBS. For actin visualization, phalloidin Alexa 488 or Alexa 568 in 1× PBS (15 μL of stock phalloidin per 200 μL of PBS) was used for 2 hr at room temperature. For immunofluorescence experiments, cells were blocked in 5% bovine serum albumin (BSA) in PBS for 20 min, followed by antibody incubations.
For visualizing titin and myomesin, a live-cell extraction was performed to remove cytoplasmic background (Svitkina, 2009). A cytoskeleton-stabilizing live-cell extraction buffer was made fresh containing 2 mL of stock solution (500 mM 1,4-piperazinediethanesulfonic acid, 25 mM ethylene glycol tetra acetic acid, 25 mM MgCl2), 4 mL of 10% polyoxyethylene glycol (PEG; 35,000 molecular weight), 4 ml H2O, and 100 μL of Triton X-100, 10 μM paclitaxel, and 10 μM phalloidin. Cells were treated with this extraction buffer for 1 min, followed by a 1 min wash with wash buffer (extraction buffer without PEG or Triton X-100). Cells were then fixed with 4% PFA for 20 min, followed by antibody labeling. VectaShield with DAPI (H-1200, Vector Laboratories Inc, Burlingame, CA) was used for mounting.
Protein knockdown
Knockdowns for MYOM were performed using single siRNAs from GE Dharmacon: one to the UTR of MYOM and one to the coding sequence (CDS). Knockdowns for MYH6 were performed using one of two siRNAs for MYH6: one to the UTR and one to the CDS purchased from GE Dharmacon. Knockdowns for MYH7 were performed using one of two siRNA pools: one of two siRNAs and one of four. Experiments were performed in 96-well culture plates, using the Lipofectamine RNAiMAX reagent and instructions provided by the manufacturer (Thermo Fisher, LMRNA015). Following knockdown, cells were replated onto glass substrates for 24 hr and fixed for immunofluorescence or lysed for western blotting.
Western blotting
Gel samples were prepared by mixing cell lysates with LDS sample buffer (Life Technologies, NP0007) and sample reducing buffer (Life Technologies, NP0009) and boiled at 95°C for 5 min. Samples were resolved on Bolt 4–12% gradient Bis-Tris gels (Life Technologies, NW04120BOX). Protein bands were blotted onto a nylon membrane (Millipore). Blots were blocked using 5% nonfat dry milk (NFDM, Research Products International, Mt. Prospect, IL, M17200) in Tris-buffered saline with Tween-20 (TBST, 10× TBS from Corning 46-012CM, Tween-20 from Sigma P7949). Antibody incubations were also performed in 5% NFDM in TBST. Blots were developed using the Immobilon Chemiluminescence Kit (Millipore, WBKLS0500).
Fluorescence microscopy
Imaging was performed using either Spinning Disk confocal microscopy or instant Structured Illumination Microscopy (iSIM). The spinning disk images were taken on a Nikon Spinning Disk confocal microscope equipped with Apo TIRF Oil 100 × 1.49 NA objective and a Photometrics Prime 95B cMOS monochrome camera, provided by the Nikon Center of Excellence, Vanderbilt University. Images were deconvolved post-acquisition using the FIJI Microvolution software plugin (Microvolution, Cupertino, CA). iSIM imaging was performed with a Visitech iSIM using a Nikon SR HP Apo TIRF 100× oil immersion objective (model number MRD01997) at 1× zoom with NA = 1.49. Images were captured using a Hamamatsu ORCA-Fusion Digital CMOS camera (model C14440-20UP) with a 0.1 µm axial step size. Images were deconvolved using Microvolution software (Cupertino, CA) installed in FIJI (Fiji Is Just ImageJ) over 20 iterations.
Platinum replica transmission EM of live-cell extracted cells
Adherent plasma membranes from cultured cardiomyocytes grown on glass coverslips were detergent extracted. Cells were treated with extraction buffer (2 mL stock buffer [5× Stock Buffer: 500 mM 1,4-piperazinediethanesulfonic acid, 25 mM ethylene glycol tetraacetic acid, 25 mM MgCl2, pH'd and kept at 4°C], 4 mL 10% PEG [35,000 MW], 4 mL milliQ H2O, 100 uL of TritonX-100, 10 uM nocodazole, and 10 uM phalloidin) for 30 min, followed by a 1 min wash with wash buffer (2 mL stock buffer, 8 mL milliQ H2O, 10 uM nocodazole, 10 uM phalloidin), followed by fixation (2% PFA, 2% glutaraldehyde) for 20 min. Extracted cells were further sequentially treated with 0.5% OsO4, 1% tannic acid, and 1% uranyl acetate before graded ethanol dehydration and hexamethyldisilazane (HMDS) substitution (LFG Distribution, France). Dried samples were then rotary shadowed with 2 nm of platinum (sputtering) and 4–6 nm of carbon (carbon thread evaporation) using an ACE600 metal coater (Leica Microsystems, Germany). The resultant platinum replica was floated off the glass with hydrofluoric acid (5%), washed several times on distilled water, and picked up on 200 mesh formvar/carbon-coated EM grids. The grids were mounted in a eucentric side-entry goniometer stage of a transmission electron microscope operated at 120 kV (JEOL, Japan), and images were recorded with a Xarosa digital camera (EM-SIS, Germany). Images were processed in Adobe Photoshop to adjust brightness and contrast and presented in inverted contrast.
Statistical analyses
Analyses comparing three groups (one control and two treatment groups: Figures 3C–G–5B–F—8C–G, Figure 3—figure supplement 1A–L, Figure 3—figure supplement 2A–L, Figure 3—figure supplement 3A–L, Figure 5—figure supplement 1A–N, Figure 6—figure supplement 1A–G, Figure 7—figure supplement 2A–N, Figure 7—figure supplement 3A–G, Figure 7—figure supplement 6A–G, Figure 8—figure supplement 1A–L, Figure 8—figure supplement 2A–N) were calculated using a one-way ANOVA. If significant, a post-hoc Tukey test (Figures 3C–F–5D—8D–G, Figure 3—figure supplement 2C, D, F, G, K, Figure 3—figure supplement 3A–D, Figure 5—figure supplement 1C, D, K, M, Figure 6—figure supplement 1A–D, Figure 7—figure supplement 2N, Figure 7—figure supplement 3F, G, Figure 7—figure supplement 5E, F, N, Figure 7—figure supplement 6D, F, G, Figure 8—figure supplement 1A, D, K, Figure 8—figure supplement 2F, I, L, N.) was done. Analyses comparing two groups were calculated using a two-tailed unpaired Student’s t-test (Figure 8A–L, S11A–L, Figure 7—figure supplement 4A-L). Graphs were made using SuperPlots (Lord et al., 2020).
Additional informationCompeting interests
No competing interests declared
Author contributions
Conceptualization, Data curation, Software, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing – original draft, Writing – review and editing
Funding acquisition, Investigation, Writing – review and editing
Funding acquisition, Investigation, Methodology
Conceptualization, Funding acquisition, Writing – review and editing
Conceptualization, Funding acquisition, Writing – review and editing
Investigation, Writing – review and editing
Investigation, Writing – review and editing
Conceptualization, Funding acquisition, Methodology, Writing – original draft, Writing – review and editing
Additional filesData availability
SarcApp and yoU-Net software can be found at Github. https://github.com/abbieneininger/sarcApp, (copy archived at Abbieneininger, 2023).
Acknowledgements
We would like to thank the instructors, TAs, and fellow classmates at the Marine Biosciences Laboratory course Deep Learning for Biological Microscopy, especially Jan Funke (Janelia), Dagmar Kainmueller (MDC Berlin), Zachary Whiddon (University of Louisville), and William Patton (Janelia) for assistance on early versions of yoU-Net. We would also like to thank Bryan Millis at the Vanderbilt Biophotonics Center and Kari Seedle at the Vanderbilt Nikon Center for Excellence and the Vanderbilt Center for Imaging Shared Resources (CISR) for experimental and imaging assistance. We thank Vanderbilt’s Program in Developmental Biology, Microtubules and Motors Club and Molecular Biophysics Training Program for project feedback and discussion. This work was supported by Vanderbilt University T32 5T32HD007502-25 to ACN and ZCS, Vanderbilt University T32 5T32GM008320-32 to JBH, American Heart Predoctoral Fellowship 836090 to JBH and 18PRE33960551 to NT, NIH NIGMS R35 GM125028 to DTB, Vanderbilt University R25 5R25GM062549-18 and American Heart Predoctoral Fellowship 1070985 to ZCS, and F31 HL136081 to AMF.
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Basic to Translational Science372874010.1016/j.jacbts.2018.08.00830623132YinXXSunLFuYLuRZhangY2022U-Net-Based Medical Image SegmentationJournal of Healthcare Engineering2022418978110.1155/2022/41897813546366010.7554/eLife.87065.3.sa0eLife assessmentDuboisNicole CReviewing EditorIcahn School of Medicine at Mount SinaiUnited StatesSolidUseful
This manuscript describes a useful tool for quantitative assessment of sarcomere structures in healthy and perturbed cardiomyocytes grown in vitro. The work is solid, and the methods, data and analyses broadly support the claims with only minor weaknesses. The tool will be relevant to biologists working on and interested in obtaining quantitative information on sarcomere structure, function and development.
This manuscript by Neininger-Castro and colleagues presents a novel automatic image analysis method for assessing sarcomeres, the basic units of myofibrils and validates this tool in a couple of experimental approaches that interfere with sarcomere assembly in iPSC-cardiomyocytes (iPSC-CM).
Automatic quantification of sarcomeres is definitely something that is useful to the field. I am surprised that there is no reference in the manuscript to SarcTrack, published by Toepfer and colleagues in 2019 (PMID 30700234), which has exactly the same purpose. The advantage of the image analysis software presented in the current manuscript appears to me to be that it can cover both mature sarcomeres and nascent sarcomeres in premyofibrils effectively.
Neininger-Castro et al report on their original study entitled "Independent regulation of Z-lines and M-lines during sarcomere assembly in cardiac myocytes revealed by the automatic image analysis software sarcApp", In this study, the research team developed two software, yoU-Net and sarcApp, that provide new binarization and sarcomere quantification methods. The authors further utilized human induced pluripotent stem cell-derived cardiomyocytes (hiCMs) as their model to verify their software by staining multiple sarcomeric components with and without the treatment of Blebbistatin, a known myosin II activity inhibitor. With the treatment of different Blebbistatin concentrations, the morphology of sarcomeric proteins was disturbed. These disrupted sarcomeric structures were further quantified using sarcApp and the quantification data supported the phenotype. The authors further investigated the roles of muscle myosins in sarcomere assembly by knocking down MYH6, MYH7, or MYOM in hiCMs. The knockdown of these genes did not affect Z-line assembly yet the knockdown of MYOM affected M-line assembly. The authors demonstrated that different muscle myosins participate in sarcomere assembly in different manners.
Neininger-Castro and colleagues developed software tools for the quantification of sarcomeres and sarcomere-precursor features in immunostained human induced pluripotent stem cell-derived cardiac myocytes (hiCMs). In the first part they used a deep-learning- based model called a U-Net to construct and train a network for binarization of immunostained cardiomyocyte images. They also wrote graphical user interface (GUI) software that will assist other labs to use this approach and made it publicly available. They did not compare their approach to existing ones, but example from one image suggests their binarization tool outperforms Otsu thresholding binarization.
In the second part they developed a software tool called sarcApp that classifies sarcomere structures in the binarized image as a Z-Line or Z-Body and assigns each to either a myofibril or to stress fibers. The tools can then automatically count and measure multiple features (33 per cell and 24 per myofibril) and report them on a per-cell, per-myofibril, and per- stress fiber basis.
To test the tools they used Blebbistatin to inhibit sarcomere assembly and showed that the sarcApp tool could capture changes in multiple features such as fewer myofibrils, fewer Z-Lines, decreased myofibril persistence, decreased Z-Line length and altered myofibril orientation in the Blebbistatin treated cells. With some changes the tool was also shown to quantify sarcomeres in titin and myomesin stained cardiomyocytes.
Finally they used sarcApp to quantify the changes in sarcomere assembly after siRNA mediated knockout of MYH7, MYH7, or MYOM. The analysis indicates that neither MYH6 nor MYH7 knockdown perturbed the assembly of Z- or M-lines, and that knockdown of MYOM perturbed the A-band/M-Line but not the Z-Line assembly according to features captured by the sarcApp tool.
Overall the authors developed and made publicly available an excellent software tool that will be very useful for labs that are interested in studying sarcomere assembly. Multiple features that are difficult to measure or count manually can be automatically measured by the software quickly and accurately.
There are however some remaining questions about these tools:
1. The binarization tool which is tailored to sarcomere image binarization appears promising but was not systematically compared with existing approaches. Example from one cell suggests it outperforms Otsu's binarization approach.
2. How robust is the tool? The tool was tested on images from one type of cardiomyocytes (hiCMs) taken from one lab using Nikon Spinning Disk confocal microscope equipped with Apo TIRF Oil 100X 1.49 NA objective or instant Structured Illumination Microscopy (iSIM), using deconvolution (Microvolution software) and in a specific magnification. It remains to be seen whether the tool would be equally effective with images taken with other microscopy systems, with other cardiomyocytes (chick or neonatal rat), with different magnifications, live imaging, etc. The authors state that this approach is also useful in other situations, but the data is not included in this manuscript.
3. The tool was developed for evaluation of sarcomere assembly. The authors show that for this application it can detect the perturbation by Blebbistatin, or knockdown of sarcomeric genes. It remains to be seen if this tool is also useful for assessment of sarcomere structure for other questions beside sarcomere assembly and in other sarcomere pathologies.
The following is the authors’ response to the original reviews.
Reviewer #1 (Public Review):
This manuscript by Neininger-Castro and colleagues presents a novel automatic image analysis method for assessing sarcomeres, the basic units of myofibrils and validates this tool in a couple of experimental approaches that interfere with sarcomere assembly in iPSCcardiomyocytes (iPSC-CM).
Automatic quantification of sarcomeres is definitely something that is useful to the field. I am surprised that there is no reference in the manuscript to SarcTrack, published by Toepfer and colleagues in 2019 (PMID 30700234), which has exactly the same purpose. The advantage of the image analysis software presented in the current manuscript appears to me to be that it can cover both mature sarcomeres and nascent sarcomeres in premyofibrils effectively.
We whole-heartedly disagree that SarcTrack has the exact same purpose as sarcApp. sarcApp measures more than the frequency of actinin2 images, and can measure real-space quantifications of actinin, myomesin, and titin, which has not been done before in this way. However, SarcTrack is an interesting method that we hope many researchers find helpful in their research. SarcTrack is a particle tracker that outputs the dimensions of the objects found, but does not distinguish between Z-Lines and other actinin2-positive structures (Z-Bodies, adhesions). It also does not group these structures into higher order structures such as myofibrils and muscle stress fibers.
When going through the manuscript there were a few issues that should be addressed in a revised version of the manuscript:
1. I am a bit puzzled that they took 1.4 um length as a cutoff length for a mature A-band in their quantifications, since the consensus in the field for thick filament length seems to be 1.6 um?
We use 1.4 µm as a cutoff length for the length of a Z-Line rather than the A-Band. We believe the reviewer is referring to the width of the A-Band perpendicular to the Z-lines, which is indeed 1.6 µm. However, we are referring to the length of the Z-Lines, which can span anywhere from1.4 µm to up to 10 or more µm. Thank you for allowing us to make the clarification.
1. When doing the knockdown for alpha and beta-myosin heavy chain, respectively, why did they not also do a Western blot for the "other" isoform as well (Figure 7)? We know that iPSCCM express a mixture, so the relatively mild phenotype that they observe in single knockdown experiments may well be due to concomitant upregulation of the expression of the other isoform. In my point of view this should be checked.
It is likely that in the single knockdown experiments the other isoform is upregulated, which is why we were careful in stating that neither muscle myosin alone is required for sarcomere formation. We do agree this would be an interesting experiment to check beyond the scope of this manuscript.
1. There seems to be a disconnect between the images for myomesin knockdown shown in Figure 8H and the quantification shown in Figure 8I, which makes me wonder whether the image shown in H middle (MYOM1 (1) KD), where the beta-myosin doublets do not seem to be much affected is really representative?
The image shown in the middle of H is representative of the mean length of beta-myosin doublets in MYOM1 (1) KD hiCMs. While the beta-myosin doublets are still present and organized, they are significantly shorter. In the zoomed out image, you can appreciate much shorter arrays of beta-myosin doublets that, while extending across the entire cell, are thinner than control cells.
Reviewer #2 (Public Review):
Neininger-Castro et al report on their original study entitled "Independent regulation of Z-lines and M-lines during sarcomere assembly in cardiac myocytes revealed by the automatic image analysis software sarcApp", In this study, the research team developed two software, yoU-Net and sarcApp, that provide new binarization and sarcomere quantification methods. The authors further utilized human induced pluripotent stem cell-derived cardiomyocytes (hiCMs) as their model to verify their software by staining multiple sarcomeric components with and without the treatment of Blebbistatin, a known myosin II activity inhibitor. With the treatment of different Blebbistatin concentrations, the morphology of sarcomeric proteins was disturbed. These disrupted sarcomeric structures were further quantified using sarcApp and the quantification data supported the phenotype. The authors further investigated the roles of muscle myosins in sarcomere assembly by knocking down MYH6, MYH7, or MYOM in hiCMs. The knockdown of these genes did not affect Z-line assembly yet the knockdown of MYOM affected M-line assembly. The authors demonstrated that different muscle myosins participate in sarcomere assembly in different manners.
Reviewer #3 (Public Review):
Neininger-Castro and colleagues developed software tools for the quantification of sarcomeres and sarcomere-precursor features in immunostained human induced pluripotent stem cellderived cardiac myocytes (hiCMs). In the first part they used a deep-learning- based model called a U-Net to construct and train a network for binarization of immunostained cardiomyocyte images. They also wrote graphical user interface (GUI) software that will assist other labs in using this approach and made it publicly available. They did not compare their approach to existing ones, but an example from one image suggests their binarization tool outperforms Otsu thresholding binarization.
In the second part they developed a software tool called sarcApp that classifies sarcomere structures in the binarized image as a Z-Line or Z-Body and assigns each to either a myofibril or to stress fibers. The tools can then automatically count and measure multiple features (33 per cell and 24 per myofibril) and report them on a per-cell, per-myofibril, and per- stress fiber basis.
To test the tools they used Blebbistatin to inhibit sarcomere assembly and showed that the sarcApp tool could capture changes in multiple features such as fewer myofibrils, fewer Z-Lines, decreased myofibril persistence, decreased Z-Line length and altered myofibril orientation in the Blebbistatin treated cells. With some changes the tool was also shown to quantify sarcomeres in titin and myomesin stained cardiomyocytes.
Finally they used sarcApp to quantify the changes in sarcomere assembly after siRNA mediated knockout of MYH7, MYH7, or MYOM. The analysis indicates that neither MYH6 nor MYH7 knockdown perturbed the assembly of Z- or M-lines, and that knockdown of MYOM perturbed the A-band/M-Line but not the Z-Line assembly according to features captured by the sarcApp tool.
Overall the authors developed and made publicly available an excellent software tool that will be very useful for labs that are interested in studying sarcomere assembly. Multiple features that are difficult to measure or count manually can be automatically measured by the software quickly and accurately.
There are however some remaining questions about these tools:
1. The binarization tool which is tailored to sarcomere image binarization appears promising but was not systematically compared with existing approaches.
We compared it with the existing approach we used previously in the lab, which was Otsu’s method for binarization. We are not aware of several other binarization approaches to compare to, other than using other machine learning techniques that are less advanced than a U-Net, the current standard in image-to-image translation.
1. How robust is the tool? The tool was tested on images from one type of cardiomyocytes(hiCMs) taken from one lab using Nikon Spinning Disk confocal microscope equipped with Apo TIRF Oil 100X 1.49 NA objective or instant Structured Illumination Microscopy (iSIM), using deconvolution (Microvolution software) and in a specific magnification. It remains to be seen whether the tool would be equally effective with images taken with other microscopy systems, with other cardiomyocytes (chick or neonatal rat), with different magnifications, live imaging, etc.
We tested the software with several magnifications, with live imaging, and with other tissues. We did not include the information in the manuscript because the data we tested the software with is for future manuscripts studying different aspects of sarcomere formation and maintenance. sarcApp reliably identifies Z-Lines and sarcomeres with deconvolved widefield fluorescence images of hiCMs and frozen human tissue, and are currently using it to measure zebrafish data for another study. Further, it works for live imaging with an actinin2-GFP (or similar) label. For the titin quantification, we would recommend using only 60-100X magnification, as the titin structures (doublets and rings) are not resolvable at lower magnifications.
1. The tool was developed for evaluation of sarcomere assembly. The authors show that for this application it can detect the perturbation by Blebbistatin, or knockdown of sarcomeric genes. It remains to be seen if this tool is also useful for assessment of sarcomere structure for other questions beside sarcomere assembly and in other sarcomere pathologies.
While this is beyond the scope of this specific methods paper, we welcome other researchers to use our software for other questions in other pathologies. We are currently doing the same for other manuscripts from our lab.
Reviewer #1 (Recommendations For The Authors):
1)"alpha-actinin..., which border the sarcomeric contractile machinery (thin and thick filaments); Z-lines do NOT border thick filaments in a relaxed sarcomere
We have removed “(thin and thick filaments)” from the text.
1. myomesin targeting siRNAs (gene name MYOM): there are actually three genes encoding for myomesin family members, specify, which one was targeted (I am assuming MYOM1).
Thank you for the clarification: we do target MYOM1
1. I am not surprised that they found not many mature Z-lines in the absence of both sarcomeric myosins; a similar codependence of assembly of mature Z-discs and the presence of functional thick filaments was previously shown by Geach and colleagues in 2015 (PMID 25845369)
Thank you for sharing this manuscript: we have added a reference to it in our study.
Reviewer #2 (Recommendations For The Authors):
This work offers the possibility to gain more insights into the process of sarcomere assembly through the advancement in sarcomeric or myofibril structure analyses. However, some clarifications are needed from the authors, please see below for the comments.
1. It is recommended that the authors include the time points for replating and harvesting hiCMs. After replating, the cardiomyocytes require at least three to four days for sarcomeric structures to reform. If the hiCMs were fixed before sarcomere assembly had completed, the staining of sarcomeric proteins including ACTN2 and titin could be compromised and it is difficult to tell if the phenotypes observed were consequences of drug treatments or knockdown of sarcomeric genes or simply because the replating hiCMs were fixed before their sarcomeric structures had fully regrown. It is also recommended that the authors replate hiCMs at a fixed time point to avoid discrepancies in the data.
Cardiomyocytes do not require three to four days for sarcomeric structures to re-form, and indeed only require 24 hours, with the first sarcomeres typically appearing at ~6 hours. We and others have published several studies demonstrating this (Fenix et al., eLIfe 2018, Taneja, Neininger and Burnette MBoC 2020, Chen et al. Nature Methods, 2022). While sarcomeres continue to develop and turn over after this time, our lab is interested in the beginning steps of sarcomerogenesis rather than the turnover of mature structures.
1. The sarcApp automatically identifies Z-lines and Z-bodies; however, is there an option for the users to set their own thresholds? Some users may select different criterions when quantifying sarcomeres. Moreover, the Z-lines and Z-bodies identified by the software are not always accurate. Can the users modify the list manually in an unbiased way. If this function is not available, the authors may consider adding this function to their software. sarcApp measures Zline and Z-bodies length but does not measure Z-line and Z-bodies width, but sometimes it is also necessary to measure the width.
Absolutely, users can modify the thresholds to identify Z-Lines and Z-Bodies. There is not a way for users to modify the list in an unbiased way per se, as editing the list of Z-Lines and Z-Bodies based on non-mathematical measurements is inherently biased, but the user is free to add in other Z-Lines and Z-Bodies as they wish. In this context, “manually” and “unbiased” is mutually exclusive.
1. It is recommended that the authors include the original images beside the sarcomeric structures identified by sarcApp (Figure 2A, 2C, 4C-F and more). It would be easier to compare the original Z-lines and Z-bodies with those identified by the software.
We have added these in Author response image 1.
Uncropped images and merges from Figures 2, 4 and 6, respectively.
1. The M-line length quantification data in Figure 3G, 5F, and 6H showed different colored-dots labeling n1 to n3, but the authors did not discuss the significance of these symbols.
We are not sure what the reviewer means by this statement: there is no significance of the different colored dots other than to mark the biological replicate shown. These graphs were created using SuperPlots, which was not stated in the original methods. It has now been added to the Statistical Analysis section.
1. Can the authors elaborate more on the reasons why they treated Blebbistatin at concentrations of 50µM and 100µM. Previous studies showed that 25µM of Blebbistatin was sufficient to delay the transformation of cardiomyocytes (PMID 27072942). Can the authors also comment on why they selected 6 hours, 12 hours, and 24 hours post replating for drug treatment. Moreover, the drug treatment at different time points was only done on ACTN2 but not titin or myomesin.
We selected 6, 12, and 24 hours for actinin2 to show the time course of sarcomere formation and to show that sarcomeres are developed by 24 hours, as also mentioned above. We are interested in future studies of the time course of titin and myomesin over time, and are working on it in the lab.
We chose 50 and 100 µM Blebbistatin as these completely blocked sarcomere assembly whereas treatment with 25 µM did not. This manuscript is a methods paper that aims to validate sarcApp and show how it could be used. We did not intend for it to be a comprehensive study of how different concentrations of blebbistatin affects sarcomere assembly.
We are also unsure what the reviewer means by “transformation of cardiomyocytes”. The manuscript with the PMID of 27072942 does not address this issue. The paper is a “review and analyze readmission data for patients who received a continuous flow left ventricular assist device (LVAD)”. We assume the reviewer is referring to differentiation. The model system we developed and published in eLife in 2018 does not use differentiating iPSC cardiac myocytes. The hiCMs we use are terminally differentiated but still immature, as they are more transcriptionally similar to primary fetal myocytes. As such, they do not maintain their sarcomeres when they removed from the 96 well and plated onto a glass coverslip for highresolution microscopy. These assemble sarcomeres within 24 hours with the sarcomeres forming close to the dorsal membrane and then rearrange overtime (e.g., moving from the top of the cell to the bottom) (Fenix et al., eLife 2018). With that said, we do agree with the reviewer that a study of sarcomere assembly in the context of cardiac myocyte differentiation would be a fascinating direction for future studies, and we think sarcApp could facilitate such studies.
1. The authors mentioned that the myofibrils of Z-line, titin, and M-line were randomly oriented after Blebbistatin treatments. The myofibrils were randomly oriented for titin and M-line. However, the orientation of Z-line after 50µM Blebbistatin treatment was not necessarily random, only the orientation after 100µM Blebbistatin treatment was randomized. The authors might consider changing bar graph to other types of charts if the orientation was really randomized after quantification.
We find that the bar chart is the most informative to us, but users can consider other types of charts in their analyses.
1. It is recommended that the authors include images staining ACTN2 at lower magnifications (Figure 1A, 1C). With current images, it is true that yoU-Net can separate Z-lines from Z-bodies yet it is difficult to tell if yoU-Net can still distinguish Z-lines from Z-bodies with larger images or it only applies to a small portion of the image.
The yoU-Net can distinguish Z-Lines from Z-Bodies with images of any size, as image size (height vs. width in pixels) does not affect how binarization occurs. During binarization, the only pixel requirement is that the width and height are divisible by 8 (for downsampling purposes). Usually this is not the case with raw images, so the image borders are slightly cropped to make them usable. In terms of resolution, we recommend using 60X-100X objectives on confocal or superresolution data for the clearest results. We have, however, successfully binarized deconvolved widefield images at 100X as well.
1. The authors mentioned that the knockdown of MYH7 did not affect Z-lines and M-lines; however, the structures of ACTN2, myomesin, and titin appeared more organized as compared to those in control.
We agree that the sarcomeres and myofibrils look slightly more organized, and did mean to state that the knockdown did not negatively affect Z-Lines and M-Lines and have updated the manuscript to be more accurate.
1. Please provide the merge images for Fig. 4D, 4E, 6B
The merge images for Fig. 4D, 4E, and 6B are included with the original images requested above (point 3)
1. In the text, they described" "antibodies to the titin I-band localize to both MSFs and sarcomeres in hiCMs (Figure 4A). Titin forms ring-like structures around the Z-Bodies of MSFs that are closer to the apparent sarcomere transition point (Figure 4A)" However, based on the antibody information they provided, it is not explicitly recognized for N-or C-terminus TITIN. Please provide TTN N-terminus or TTN-C terminus co-stainings with ACTN2 antibody to understand which part of TTN together with ACTN2 forms a Z-Body.
The TTN antibody is an N-terminal antibody localizing to the I-Band region of sarcomeres. We agree with the reviewer that a more thorough study of titin will be of interest and we are currently undertaking such a study. However, this is a methods paper presenting a tool. While some of the data we present does point to mechanistic hypotheses, it is beyond the scope of this study to fully characterize titin during sarcomere assembly.
1. TITIN doublet was used to indicate a sarcomere in Fig. 4C-D. Moreover, they also used another combination (myomesin and F-ACTIN) to label a sarcomere in Fig. 6D. Can they compare the difference between these two methods or by using these two methods (TITIN doublet) and (myomesin and F-ACTIN), how is the average length of sarcomere? Will the sarcomere length be the same?
We noted in the manuscript that due to the organization of titin doublets (wrapping around the ends of Z-Lines) that the average titin doublet will be approximately 0.3 um longer than the ZLine. We did not expect to see a difference in lengths of myomesin M-Lines and mature actinin2 Z-Lines and indeed do not see major differences in the average lengths (between 2.0 and 2.5 um in 24 hour control cells)
1. They used siRNA method to knockdown MYH6, MYH7 and MYOM and concluded that the knockdown of these genes did not affect the Z-line assembly. Even though they showed very nice knockdown efficiency of these proteins, they should (1) co-stain MYH6/TITIN/actinin2 and MYH6/ myomesin /actinin2 for Fig. 7C. (2) MYH7/TITIN/actinin2 and MYH7/ myomesin /actinin2 for Fig. 7I. (3) MYOM1/TITIN/actinin2 and MYOM2/TITIN/actinin2 for Fig. 8A. (4) MYH7/MYOM1 and MYH7/MYOM2 for Fig. 8H to make sure the cells they measured were truly knockdownpositive cells,
The antibodies for alpha and beta myosin are not very efficient for immunofluorescence, and work best for western blots. We decided also to choose a random subset of the cells on the dish to be sure to eliminate any risk of cherry-picking. While imaging cells on the dish, we looked only at the DAPI nuclear channel and selected 50 cells minimum per dish with only this channel, then imaged the other channels.
Minor comments:
1. Well-organized sarcomere structure on DMSO treated cells in Fig.5A and Fig. 6A, but it was disarray in Fig. S3M. Why?
Figure S3 shows hiCMs that have only been allowed to spread for 6 hours, which have not formed mature sarcomeres yet, hence the disarray.
1. Fig 1A, Fig2B: please label the name of the antibody, not the actin filament
We used phalloidin labelling here, which marks actin filaments. We have updated the figure legends to be more clear. Thank you!
1. Fig. 7I: actinin2 instead of actinin
Thank you for catching this! We have fixed it.
Reviewer #3 (Recommendations For The Authors):
Testing the app using images shot by other microscopy systems, magnifications, and cardiomyocytes from other species, as noted in the public review above, should make the app even more wildly useful.
A more formal head-to-head comparison with other approaches will be more convincing in showing the new tool is superior
I also think that a more detailed protocol for using the app will help other investigators.
The app counts and measures many features, but it is not always clear how and using what algorithm these are measured. Including these details in a protocol or even as comments in the code will be very helpful for others.
The protocol found on the public GitHub for the app will help other investigators to download, use, and understand the application. We have received contact from researchers who have been able to use the application without assistance from us, which is a good sign that the application is user-friendly and that the online protocol is sufficient.